Control of the electrostatic potential of nanoparticles

ABSTRACT

The present technology is directed to the nanoparticles for use as molecular environmental sensors. The nanoparticles comprise a photoluminescence core and a plurality of ligands bound to the core and forming a quencher permeable ligand shell. The ligands comprise a reactive or charged moiety capable of being modulated between a first stand and a second state, and the proportion of ligands in each state determine the permeability of the ligand shell to a photoluminescence quencher.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application claims benefit of priority to U.S. ProvisionalApplication No. 62/523,980, filed 23 Jun. 2017, the content of which isincorporated herein by reference it its entirety.

STATEMENT OF GOVERNMENT SUPPORT

This invention was made with government support under R21 GM127919awarded by the National Institutes of Health (NIH) and CHE 1664184, CHE1400596, and DMR 1121262 awarded by the National Science Foundation(NSF). The government has certain rights in the invention.

FIELD OF INVENTION

The present technology is related to nanoparticles having a controllableelectrostatic potential and methods of preparation and use. Morespecifically the present technology is related to environmental sensorscomprising nanoparticles that have a permeable ligand shell composed ofreactive or charged moieties that may be modulated between states inresponse to its molecular environment.

BACKGROUND

Colloidal quantum dots (QDs), synthesized with wet chemical methods,form a class of highly versatile, solution-processable nanoscalebuilding blocks for bottom-up fabrication of hierarchical structureswith wide-spread potential applications, such as solid stateelectronics, solar cells, photocatalysts and biological tags. Theperformance of QDs in any of these applications depends on control lingtheir interactions with small molecules and ions that, for example,quench their photoluminescence (PL) through charge or energy transfer,corrode their surfaces through reactions with surface ions or ligands,or induce aggregation of particles. One method for maximizing theinteraction of QDs with specified molecules and minimizing non-specificinteractions is to use the self-assembled monolayer (SAM) that serves asthe ligand shell for the QD as a semi-permeable membrane, and,ultimately, a molecular recognition layer. An important technique tocreate selectively permeable membranes is to make them electricallycharged. The density and type of these charges control the electrostaticpotential at the membrane-solvent interface. As a result, there is aneed for new and improved methods for controlling the electrostaticpotential of nanoparticles at the surface.

SUMMARY OF THE INVENTION

Disclosed herein sensors for monitoring chemical and biological process.The sensors detect changes in the molecular environment by modulatingbetween reactive or charged states. The sensors comprise aphotoluminescence quencher and a nanoparticle. The nanoparticlecomprises a photoluminescent core and plurality of reactive ligandsbound to the core and forming a quencher permeable ligand shellsurrounding the core. Each of the reactive ligands comprise a reactivemoiety capable of being modulated between a first state and a secondstate and an anchoring group for binding the ligand to the core, and thepermeability of the ligand shell is determined by the proportion of theligands in a first state and a second state. The nanoparticle mayfurther comprise a diluent ligand, a solubilizing ligand, or both thediluent ligand and the solubilizing ligand.

In some embodiments, the reactive moiety comprises an anionic charge inthe first state and a neutral charge in the second state. In otherembodiments, the reactive moiety comprises a cationic charge in thefirst state and a neutral charge in the second state.

In some embodiments, the reactive ligand comprises a radical of formulaA-T-R. A may comprises the anchoring group and the anchoring group isselected from the group consisting of a alkylmonothiolate, aalkyldithiolate, or an alkyl trithiolate. R may comprise a reactivemoiety selected from the group consisting of a carboxyl, a hydroxyl, asulfo, a sulfhydryl, a phosphoryl, a phosphate and a conjugate basethereof or a reactive moiety selected from the group consisting of asubstituted or unsubstituted amine or alkylamine, a substituted orunsubstituted imidazole, a substituted or unsubstituted benzimidazole, asubstituted or unsubstituted pyrimidine, a substituted or unsubstitutedpurine, a substituted or unsubstituted pyridine, a substituted orunsubstituted pyrrolidine, and a conjugate acid thereof. T may comprisea tether comprising —(CH₂)_(n)— where n is an integer and n=1-15,—(CH₂)_(n)(CONH)(CH₂)_(m)— where n and m are integers and n+m=1-15, or—(OCH₂CH₂)_(n)— where n is an integer and n=1-100.

In some embodiments, the quencher comprises an anionic charge orcationic charge. In some embodiments, the quencher comprises9,10-anthraquinone-2-sulfonate, rhodamine B, or methyl propyl viologen.

In some embodiments, the sensor further comprises a flexible tether fortethering the quencher to the nanoparticle. In certain embodiments, thequencher comprises a radical of formula A-T-Q. A may comprises theanchoring group and the anchoring group is selected from the groupconsisting of a alkylmonothiolate, a alkyldithiolate, or an alkyltrithiolate. Q may comprise an aromatic moiety capable of accepting anelectron from the core. T may comprises a tether comprising —(CH₂)n—where n is an integer and n=1-15 —(CH₂)_(n)(CONH)(CH₂)_(m)— where n andm are integers and n+m=1-15, or —(OCH₂CH₂)_(n)— where n is an integerand n=1-100. I certain embodiments, the aromatic moiety comprises9,10-anthraquinone-2-sulfonate, rhodamine B, or methyl propyl viologen.

In some embodiments, the first state is a first protonation state andthe second state is a second protonation state. In other embodiments,the first state is an acetylated state and the second state is adeacetylated state.

Another aspect of the invention provides for environmental sensorcomprising a photoluminescence quencher and a nanoparticle comprising aphotoluminescent core and plurality of charged ligands bound to the coreand forming a quencher permeable ligand shell surrounding the core. Eachof the charged ligands comprises a charged moiety capable of beingmodulated between a first state and a second state and an anchoringgroup for binding the ligand to the core, and the permeability of theligand shell is determined by the proportion of the ligands in a firststate and a second state. In some embodiments, the first state is afirst protonation state and the second state is a second protonationstate. In other embodiments, the first state is an acetylated state andthe second state is a deacetylated state. In yet other embodiments, thefirst state is an ion-paired state and the second state ion-unpairedstate.

Another aspect of the invention provides for a method for environmentalmonitoring, the method comprising contacting ay of the environmentalsensors described herein with a molecular environment, irradiating thesensor, and detecting a photoluminescent signal. In some embodiments,the molecular environment comprises a reactant capable of generating aproduct and wherein the product is capable of reacting with the reactivemoiety to modulate the reactive moiety between the first state and thesecond state. The product may be H⁺, OH⁻, an acid, or a base.

In some embodiments, the molecular environment comprises a reactantcapable of generating a product and wherein the reactant is capable ofreacting with the reactive moiety to modulate the reactive moietybetween the first state and the second state. The reactant may compriseH⁺, OH⁻, an acid, a base, an acetyl, a methyl, a phosphoryl, carboxyl,hydroxyl, or amine.

In some embodiments, the molecular environment comprises an ion capableof pairing with the charged moiety to modulate the charged moietybetween the first state and the second state. The ion may comprise amember selected from the group consisting of ammonium, imidazolium,pryidinium, pyrrolidinium, phosphonium, sulfonium, and any combinationthereof.

BRIEF DESCRIPTION OF THE DRAWINGS

Non-limiting embodiments of the present invention will be described byway of example with reference to the accompanying figures, which areschematic and are not intended to be drawn to scale. In the figures,each identical or nearly identical component illustrated is typicallyrepresented by a single numeral. For purposes of clarity, not everycomponent is labeled in every figure, nor is every component of eachembodiment of the invention shown where illustration is not necessary toallow those of ordinary skill in the art to understand the invention.

FIG. 1A shows structures of the NR₄ ⁺ cations, which we add to the QDsas chloride salts: R=—CH₃, tetramethylammonium (TMA); R=—CH₂CH₃,tetraethylammonium (TEA); R=—(CH₂)₂CH₃, tetrapropylammonium (TPA); andR=—(CH₂)₃CH₃, tetrabutylammonium (TBA) cations.

FIG. 1B shows a chematic representation of NR₄ ⁺ forming ion pairs withthe cathoxylate tail groups of 6-mercaptohexanoate (MHA) ligands,thereby enhancing the permeability of this anionic ligand shell toward9,10-anthraquinone-2-sulfonate (AQ), the electron acceptor. The positiveand negative charge centers are highlighted in red and blue,respectively. We added AQ to the QDs as a sodium salt.

FIG. 2A shows the fraction of emissive PbS QDs (2.63 μM) in the ensemblethat remain emissive upon addition of 1000 equiv. of AQ quencher(PL/PL₀) vs. the molar equivalents of NR₄ ⁺ added to the CSD sample.Three measurements on separately prepared samples were averaged toobtain each data point, and the error bars are the standard deviation ofthose measurements. The sizes in the legend are the hydrodynamic radii,r, of the NR₄ ⁺ cations, measured by diffusion-ordered NMR spectroscopy(DOSY) in D₂O. Traces are TMA, TEA, TPA, and TBA from top to bottom.

FIG. 2B shows normalized kinetic traces extracted (at the ground statebleach minimum, 1014 nm) from the nanosecond-to-microsecond TA spectrumof a 6.58 μM sample of MHA-capped PbS QDs, mixed with 1000 equiv. of AQand no NR₄ ⁺ cations (black), 1000 equiv. of AQ and 50,000 equiv. TMA(red), or 1000 equiv. of AQ and 50,000 equiv. IBA (blue). The solidlines are fits using a sum of two exponentials convoluted with aninstrument response function. Traces are PbS+AQ+TBA, PbS+AQ+TMA, and PbSAQ from top to bottom.

FIG. 3 shows PL/PL₀ of a 2.63 μM solution of MHA-capped PbS QDs mixedwith 1000 equiv. of AQ and increasing equiv. of TBA, plotted as afunction of increasing TBA/AQ ratio (curved plot), and hydrodynamicradius of AQ calculated from DOSY and plotted as a function ofincreasing TBA/AQ ratio (linear plot). While PL/PL₀ saturates from 0.75to 0.2 drastically as TBA/AQ increases from 0 to 50, the hydrodynamicradius of AQ only increased gradually by a factor of 1.6; the two-foldincrease in hydrodynamic radius when TBA/AQ increases from 50 to 200,however, seems to have no effect on the yield of PL quenching.

FIG. 4A shows PL spectra of sample 1 (see Table 8), 2.63 μM PbS QDs eachcapped by 115 MHA ligands, mixed with increasing equivalents of AQ. Thedecreasing intensity of PL indicates a growing eT yield as we increasethe number of electron acceptors adsorbed per QD.

FIG. 4B shows plots of PL/PL₀, the fraction of originally emissive QDsthat remain emissive after addition of AQ, vs. concentration of free AQin solution. The lower the surface coverage of the negatively chargedMHA ligand on the QDs, the more efficiently AQ quenches the PL of theQDs. The PL quenching for the sample of the QDs capped with 115 MHAligands (shown in open symbols) has a contribution from collisional eTbetween freely diffusing species, in addition to the quenching withinstatic QD-AQ complexes.

FIG. 5A shows normalized kinetic traces extracted at the ground statebleach (1014 nm, inset) from the TA spectrum of a 6.58 μM sample of115-MHA capped PbS QDs mixed with 0 eq. (black), 2000 eq. (red) and 4000eq. (blue) of AQ. The solid lines in FIG. 5A are multi-exponential fitsof the traces with parameters summarized in Table 10.

FIG. 5B shows normalized kinetic traces extracted (at 1039 nm) from theTA spectrum of a 6.58 μM sample of 31 MHA-capped PbS QDs mixed with 0eq. (black), 100 eq. (red) and 200 eq. (blue) of AQ. The solid lines inFIG. 5B are multi-exponential fits of the traces with parameterssummarized in Table 10.

FIG. 6A shows full ¹H NMR spectra for Samples 1-6 in D₂O. The sharpsinglets at 8.33 ppm, 7.53 ppm, and 4.65 ppm correspond to sodiumformate (added as an internal standard for integration, see nextsection), chloroform residue, and H₂O impurity, respectively.

FIG. 6B shows ¹H NMR spectra for 13.2 μM MHA- and oleate-capped PbS QDs.

FIG. 7A shows normalized kinetic traces extracted (at 1014 nm) from theTA spectrum of a 6.58 μM sample of 115 MHA-capped PbS QDs mixed with 0eq. (black; bottom) and 4000 eq. (red; top) of AQ.

FIG. 7B shows normalized kinetic traces extracted (at 1039 nm) from theTA spectrum a 6.58 μM sample of 31 MHA-capped PbS QDs.

FIG. 8 shows a schematic Representation of Protonation of the ImidazoleTail Groups of the DHLA-His Ligand Shell of a PbS QD by Tol-SO₃H, andthe Effect of This Protonation on the Permeability of the Ligand Shellto AQ, a Negatively Charged Electron Acceptor.

FIG. 9A shows representative NMR spectra of a series of 0.132 mMmethanol-d4 solutions of DHLA-His-capped PbS QDs mixed with increasing(from black to red) molar equivalents (0-200 per QD) of Tol-SO₃H.—R=—C₁₀H₂₀NOS₂. The small peak at ˜7.3 ppm, marked by asterisk (*),corresponds to an unknown impurity in QDs introduced during the ligandexchange process.

FIG. 9B shows the chemical shift of Ha for DHLA-His ligands that arebound to the surface of the QD (black; lower) or freely diffusing insolution (red; upper), plotted vs the equiv of Tol-SO₃H added per QD.The errors bars are the standard deviation of three measurements onseparately prepared QD samples. The inset contains the same data forbound DHLA-His only, where the chemical shift is converted to the degreeof protonation.

FIG. 10A shows fraction of emissive PbS QDs (6.58 μM) within mixtureswith average protonation p=0 (black; upper), 31 (red), 64 (fucia), or92% (blue; bottom) that remain emissive upon addition of increasingmolar equivalents (0-50 per QD) of AQ. Three measurements on separatelyprepared samples were averaged to obtain each data point, and the errorbars are the standard deviation of these measurements. The inset showsrepresentative PL spectra of DHLA-His-capped PbS QDs (p=0) mixed withincreasing equiv of AQ.

FIG. 10B shows normalized kinetic traces extracted (at the ground-statebleach, 878 nm) from the ps-to-ns TA spectrum of a 13.2 μM sample ofDHLA-His-capped PbS QDs, mixed with 10 equiv of AQ (black; lower), 25equiv of Tol-SO₃H and 10 equiv of AQ (red), or 75 equiv of Tol-SO₃H and10 equiv of AQ (blue; upper), and photoexcited with a 100-μW laser at810 nm. The solid lines are global fits with fitting parameterssummarized in Table 13.

FIG. 10C shows normalized kinetic traces extracted (at the ground-statebleach, 901 nm) from the ns-to-μs TA spectra of the same sample seriesin FIG. 10B, photoexcited with a 2 mW laser at 810 nm. The solid linesare fits with fitting parameters summarized in Table 13.

FIG. 11 shows the integrated. PL intensity of a 3 mL, 10.0 SEM solutionof DHLA-His-capped PbS QDs, (red; upper), or QDs plus 10 equiv AQ(black; lower), upon alternate additions of 75 equiv of Tol-SO₃H and 75equiv of NMe₄OH, which we added in 10 μL aliquots. No acid or base wasadded at titration step “0” (all of the ligands are initiallydeprotonated upon ligand exchange), and all spectra were taken within 2.min of adding acid or base into the sample. In the black trace, threemeasurements on separately prepared samples were averaged to obtain eachdata point, and the error bars are the standard deviation of thesemeasurements.

FIG. 12 shows kinetic traces of T₂ decay of the Ha proton signal for thebroad (black; lower) and sharp (red; upper) features in the ₁NMRspectrum (see FIG. 9A) of 0.132 nM DHLA-His-capped PbS QDs, mixed with75 equiv. of Tol-SO₃H.

FIG. 13 shows the ratio between freely diffusing and surface-boundDHLA-His ligands as a function of Tol-SO₃H added.

FIG. 14 shows a time-dependent PL study on DHLA-His-capped PbS QDs mixedwith 10 equiv. AQ in ethylene glycol. The integrated PL intensitysaturates two hours after the addition of AQ.

FIG. 15 shows a ground-state absorption spectra of DHLA-His-capped PbSQDs with increasing degree of protonation in their ligand shells.

FIGS. 16A show ground-state absorption spectra of a 10.0 μM sample ofDHLA-His-capped PbS QDs, pretreated with 75 equiv. Tol-SO₃H and 5 equiv.AQ, before (black; upper) and after (red; lower) overnight illumination.

FIG. 16B show steady-state PL (FIG. 16B) spectra of a 10.0 μM sample ofDHLA-His-capped PbS QDs, pretreated with 75 equiv. Tol-SO₃H and 5 equiv.AQ, before (black; lower) and after (red; upper) overnight illumination.

FIG. 17 Cyclic voltammograms of AQ in ethylene glycol solutions ofincreasing acidity.

FIG. 18A shows a representative TA spectrum (extracted at a time delayof 2 ps) of a 13.2 μM ethylene glycol solution of DHLA-His-capped PbSQDs. The negative feature, indicated by the arrow, is the ground-statebleach.

FIG. 18B shows kinetic traces extracted (at the ground-state bleach, 871nm) from the ps-to-ns TA spectra of QDs with different degrees ofprotonation at their ligand shells. The fitting parameters are tabulatedin Table 14.

FIG. 18C shows kinetic traces extracted (at the ground-state bleach, 901nm) from the ns-to-μs TA spectra of the same sample series. The fittingparameters are tabulated in Table 14.

FIG. 19 shows time-dependent PL intensity of DHLA-His-capped PbS QDsupon addition of acid (lower) or base (upper).

FIG. 20 shows normalized PL Intensity of 10.0 μM PbS QD solutions mixedwith increasing molar equivalents of NMe₄Cl. PbS QDs (p=0) upper trace.PbS QDs (p=92%) lower trace.

FIG. 21 shows a schematic design and the mechanism of the electrostaticsensor.

FIG. 22 shows the dependence of the PL intensity of QDs with (red;lower) or without (black; upper) tethered MPV acceptors on pH of thesolution.

FIG. 23 shows the change of PL intensity of QDs with (red; lower)andwithout (black; upper) tethered MPV acceptors on the titration ofacetylating agent NHS-Ace.

FIG. 24 shows a schematic of a proton sensor. When the tethered quencheris in proximity to the core, eT occurs quenching photoluminescence (leftside). When the tethered quencher is extends outwardly from the core,the nanoparticle photoluminescences (right side)

DETAILED DESCRIPTION OF THE INVENTION

The present technology is directed to the control of the electrostaticpotential of a nanoparticle. The polarity and magnitude of thispotential dictate how the colloid will interact with its chemicalenvironment. This potential is a critical factor in regulating (i)permeation of small molecules to the inorganic surface of the particleand (ii) nonspecific adsorption of larger biomolecules to thenanoparticle, e.g., protein coronas that degrade the particle's abilityto tag or permeate a membrane. Introduction of a charged molecularquencher to the system renders the photoluminescence (PL) intensity ofthe nanoparticle sensitive to this potential and allows for monitoringthe yield of reactions, such as changes in the pH or acetylation, thatresult in a change in the charge or charge distribution of thenanoparticles.

The present technology monitors chemical and biological processes bytracing the change in the state of electrostatic charges. The generalmethodology relies on a ligand shell surrounding a photoluminescent corethat is permeable to a photoluminescence quencher. Changes in themolecular environmental result in a modulation between states forreactive or charged moieties within the ligand shell. For example, themoieties may modulate between a protonated and deprotonated state, anacetylated and a deacetylated state, or an ion-paired and ion-despairedstate. The change in the equilibrium distribution of states within theligand shell results in a change in the permeability of the ligand shellfor the quencher to come into proximity with the core. As a result, thedetectable photoluminescence will change and allow one to monitor themolecular environment in real-time in vitro and in vivo.

The environmental sensors comprise nanoparticles and photoluminescencequenchers. The nanoparticle comprises a photoluminescent core. A“photoluminescent core” is a material capable of emitting photons afterthe absorption of photons. Depending on the molecular environment, thecore may be selected to absorb photos or emit photons in a desiredoptical window. An example of an optical window is the near-infrared(NIR) window, which may otherwise be known as the biological window,between 650 to 1350 nm where photons have their maximum penetrationdepth in biological tissues.

The nanoparticle core may comprise any suitable material capable ofphotoluminescence. In some embodiments, the core comprises a transitionmetal, a basic metal, a semimetal, a nonmetal, or any combinationthereof. This includes, but is not limited to materials comprisingelements from Groups 11-16. The nanoparticle core may be crystalline oramporphous. In some embodiments, the nanoparticle core may be asemiconductor. Exemplary materials include, but are not limited to, CdS,CdSe, CdTe, PbS, PbSe, InP, InAs, CuInS₂, ZnSe, or ZnTe. Moreover, theterm “core” is not limited to a single material and, for example,encompasses core-shell nanoparticles.

The core may be of any suitable size that allows for photoluminesce. Insome embodiments, the nanoparticle has a core radius of less than 100nm, including nanoparticles having a core radius less than 50 nm, 25 nm,20, nm, 15 nm, 10, nm, 5 nm, or 2 nm. In some embodiments, thenanoparticle is a quantum dot.

By selecting the material and/or size of the core, the photon absorptionand photo emission widow by be tuned for a desired optical window.

Surrounding the core is a ligand shell. The ligand shell comprises aplurality of ligands bound to the core. The ligands comprise ananchoring group for binding the ligand to the core. The anchoring groupmay be any chemical moiety capable of associating the ligand to the coreby physisorption or chemisorption. Exemplary anchoring groups includethiols, such as monothiolate, dithiolate, or trithiolate groups, butother anchoring groups may be used.

In some embodiments, the ligands comprise a reactive moiety. A “reactivemoiety” may comprise any functional group capable of reacting to form abond between itself and a compound or ion in the molecular environmentor dissociating to break a bond to form a compound or ion. The bondforming or breaking reaction should alter the electrostatic potential ofthe ligand shell and/or the permeability of the shell to aphotoluminescence quencher. The bond forming or bond breaking reactionmay be reversible, but need not be. In some embodiments, the bondforming or breaking reaction may be reversible within a window fromabout 0° C. to about 50° C., including from about 0° C. to about 40° C.,10° C. to about 40° C., or 20° C. to about 40° C. In particular the bondforming or breaking reaction may include any reversible reaction at aphysiological temperature of a biochemical compound, such as areversible protein, lipid, or saccharide modification. Examples of suchreactions include, without limitation, protonation, deprotonation,phosphorylation, methylation, acetylation, hydrolysis, or condensation.

Suitably the reactive moiety may be able to modulate an acidic moietybetween its protonated state and its corresponding deprotonatedconjugate base. Reactions of this type can result in a neutral,protonated moiety switching to its anionically-charged, deprotonatedstate or a anionically-charge, deprotonated state switching to itsneutral, protonated state. A change in the environmental pH will resultin a change to the protonation equilibrium between the neutral,protonated state and the anionic, deprotonated state. This results in achange to the electrostatic potential of the nanoparticle and change inpermeability of the ligand shell. A decrease in the pH increases therelative proportion of moieties in the neutral, protonated state.Whereas an increase in the pH decreases the relative proportion ofmoieties in the neutral, protonated state. Exemplary reactive moieties,include, without limitation, a carboxyl, a hydroxyl, a sulfa, asulfhydryl, a phosphoryl, or a phosphate or a conjugate base thereof.

Suitably the reactive moiety may be able to modulate a basic moietybetween its deprotonated state and its corresponding conjugate acid.Reactions of this type can result in a neutral, deprotonated moietyswitching to its cationically-charged, protonated state or acationically-charged, protonated state switching to its neutral,deprotonated state. A change in the environmental pH will result in achange to the protonation equilibrium between the neutral, deprotonatedstate and the cationic, protonated state. This results in a change tothe electrostatic potential of the nanoparticle and change inpermeability of the ligand shell. A decrease in the pH increases therelative proportion of moieties in the cationic, protonated state.Whereas an increase in the pH decreases the relative proportion ofmoieties in the neutral, deprotonated state. Exemplary reactive moietiesinclude, without limitation, a substituted or unsubstituted amine oralkylamine, a substituted or unsubstituted imidazole, a substituted orunsubstituted benzimidazole, a substituted or unsubstituted pyrimidine,a substituted or unsubstituted purine, a substituted or unsubstitutedpyridine, a substituted or unsubstituted pyrrolidine, or a conjugateacid thereof. As demonstrated in the Examples that follow, theelectrostatic potential and permeability of the ligand shell to thequencher may be controlled by the protonation equilibrium of imidazolereactive moieties and amine reactive moieties.

Suitably the reactive moiety may be able to modulate a moiety associatedwith a post-translation protein modification. Various post-translationmodification are known and may include, without limitation,phosphorylation, methylation, or acetylation. Amino acid and peptidesites that often undergo post-translational modification are those thathave a functional group that can serve as a nucleophile in the reaction:the hydroxyl groups of serine, threonine, and tyrosine; the amine formsof lysine, arginine, and histidine; the thiolate anion of cysteine; thecarboxylates of aspartate and glutamate; and the N- and C-termini.Reversible phosphorylation occurs at the side chains of three aminoacids, serine, threonine and tyrosine. These amino acids have anucleophilic hydroxyl group that attacks the terminal phosphate group onthe universal phosphoryl donor adenosine triphosphate (ATP), resultingin the transfer of the phosphate group to the amino acid side chain.Methylation transfers a one-carbon methyl group to nitrogen or oxygen(N- and O-methylation, respectively) to amino acid side chains increasesthe hydrophobicity of the protein and can neutralize a negative aminoacid charge when bound to carboxylic acids. Acetylation transfers anacetyl group to an amine, such as the amine of lysine, increasing thehydrophobicity and can neutralize a positive amino acid charge of aprotonated amine.

Suitably the reactive moiety may be able to modulate a moiety associatedwith condensation or hydrolysis. Condensation reactions occur when twoor more reactants yield a single main product with accompanyingformation of water. Examples include the reaction of carboxyl groupswith hydroxyl groups to form esters or amines to form amides, increasingthe hydrophobicity. Hydrolysis reactions cleave a reactant with theconsumption of water to yield two products. Examples include thereaction of esters or amides with water forming a carboxyl group and ahydroxyl or amine, respectively.

In some embodiments, the ligands comprise a charged moiety. A “chargedmoiety” may comprise any functional group capable of having a persistentanionic or cationic charge in the molecular environment. The presence ofions having an opposition charge as the charged moiety may result in ionpairs that modulate the electrostatic potential of the ligand shelland/or the permeability of the shell to a photoluminescence quencher. Asdemonstrated in the Examples that follow, the electrostatic potentialand permeability of the ligand shell to the quencher may be controlledby the ion-pairing of charged carboxylate moieties withtetraalkylamines.

The ligand may have any suitable tether connecting the anchoring groupand the reactive or charged moiety. The tether may be linear orbranched, substituted or unsubstituted C₁-C₁₅ alkyl, an oligoethyleneglycol having 1, 2, 3, 4, or 5 ethylene glycol monomers, or polyethyleneglycol having 5 to 100 monomers, e.g., PEG-200, PEG-300, PEG-400,PEG-600, PEG-1000, PEG-1500, or PEG-2000. In some embodiments, theC₁-C₁₅ alkyl comprises —(CH₂)_(n)— where n is greater than or equal to 1and less than or equal to 15 or —(OCH₂CH)_(n)— where n is an integer andgreater than or equal to 1 and less than or equal to 100.

In some embodiments, the tether comprises a coupling moiety that allowsfor the modular coupling of the anchor group to a variety of reactive orcharged moieties. In some embodiments, the coupling moiety is an amideformed from the reaction of a carboxyl acid and a amine, but otherchemistries are suitable for coupling such reactions of amines withNHS-esters, isocyanates, isothiocyantes, or benzoyl fluorides; thiolateswith maleimides, iodoacetamides, 2-thiopyridine, or3-arylproplolonitrile; or azides with alkynes or other clickchemistries. In some embodiments, the tether comprises—(CH₂)_(n)(CONH)(CH₂)_(m)— where n and m are integers and n+m is greaterthan or equal to 1 and less than or equal to 15.

The ligand shell may also comprise a diluent ligand and/or asolubillizing ligand. For some applications, covering the nanoparticlecore entirely with ligands comprising a reactive or charged moiety maybe undesirable. A “diluent ligand” is a ligand that allows for thecontrolling the number of ligands comprising a reactive or charge moietyselected for the application of interest within the ligand shell,resulting in a mixed adlayer of ligands. The diluent ligand may compriseany of the anchoring groups, tethers, or coupling groups describedabove. As described in the Examples that follow, the number ofcarboxylate terminated charged groups may be controlled within theligand shell by the inclusion of hydroxyl terminated ligands.

A “solubilizing ligand” is a ligand selected to improve the solubilityof the nanoparticle in the molecular environment. The solubilizingligand comprises one or more functional groups capable of improvingsolubility. In the case of an aqueous environment, the solubilizingligand may comprise polar functional groups or a water soluble polymeror oligomer such as polyethylene glycol.

The relative number of ligands of different types in the ligand may bevaried over wide ranges depending on the application of interest. Theligand shell may comprise between about 20% (as a number percentage ofthe total number of ligands) and about 100% of the reactive ligand orthe charged ligand. In certain embodiments, the ligand shell comprisesbetween about 50% to about 95% or about 50% to about 90%. The ligandshell may comprise between about 0% and about 80% of the diluent ligand.In certain embodiments, the ligand shell comprises between about 5% andabout 50% diluent ligand or about 10% to about 50% of the diluentligand. The ligand shell may comprises between about 0% and about 25% ofthe solubilizing ligand. In certain embodiments, the ligand shellcomprises between about 1% and 10% or about 1% and about 5% of thesolubilizing ligand.

The environmental sensors further comprise a photoluminescence quencher.A “photoluminescence quencher” is any molecule or functional groupcapable of inhibiting the photoluminescence of the core. Inhibition ofthe photoluminescence may occur by any suitable mechanism, includingelectron transfer. Because the probability of electron transfer isdependent on the separation distance between the quencher and the core,modulation of the permeability of the ligand shell will modulate thequenching of the photoluminescence. Many molecules or functional groupsare suitable to accept an electron from the photoluminescent core toquench photoluminescence, including, without limitation, aromaticcompounds having one or more ring structures. The quenchers may becharged, but need not be. Examples of such quenchers used in this workinclude, 9,10-anthraquinone-2-sulfone, methyl propyl viologen, andrhodamine B.

The quencher may be optionally tethered to the photoluminescent core.The tether should be flexible to allow the relative distance between thequencher and the core to vary so that the amount of photoluminescenceinhibition can vary in response to changes in the molecular environment.An advantage of tether quenchers is that the response time of a tetheredquencher can potentially bypass the limit of diffusion (which is on theorder of tens of nanosecond to microsecond). The tether may comprise anyof the anchoring groups or coupling groups described above for attachingthe quencher moiety to the core.

The sensors described herein may be used to monitor in real-time themolecular environment. Methods of using the sensors for monitoring amolecular environment comprise contacting the sensor with a molecularenvironment, irradiating the sensor in contact with the molecularenvironment, and detecting a photoluminescent signal. The sensor withthe molecular environment may be contacted by any suitable means,including, without limitation, by preparing a solution or colloidalsystem comprising the sensor or by wetting a surface having the sensorattached thereto. For a dynamically evolving molecular environment, thephotoluminescent signal may change depending of the interaction ofmolecular species with the reactive or charged ligands present in theligand shell of the nanoparticle.

The molecular environment may comprise an ion capable of paring with acharge moiety to modulate the charged moiety between an ion-paired andion-unpaired state or an ion or compound capable of reacting with areactive moiety. The ion should be oppositely charged than the surfacedistal to the core of the nanoparticle. Exemplary cationic organic ionsinclude ammonium, imidazolium, pryidinium, pyrrolidinium, phosphonium,and sulfonium. One or more of these organic ions may be paired with thenanoparticle. For aqueous phase applications, it is preferably for theorganic ion to be at least partially soluble in water. In some cases theorganic ion comprises a C₁₋₄ substituent, including C₁₋₄ alkylsubstituents such as methyl, ethyl, propyl, and butyl substituents.Although all of the substituents of the organic ion may be the same, theorganic ion may comprise combinations of different substituents. Inparticular embodiments, the organic ion is tetramethylammonium,tetraethylammonium, tetrapropylammonium, tetrabutylammonium, or anycombination thereof.

In some cases, the dynamically evolving molecular environment involvesthe formation of a reaction product. In cases where the reactive ligandin the ligand shell is capable of reversibly or irreversibly reactingwith the product to modulate the permeability of the ligand shell, thesensor is capable of directly detecting the formation of the product. Asthe reaction progresses and product is being formed, the higherconcentration of product will change the reaction equilibrium betweenthe first and second states. Depending on how the permeability of theligand shell is modulated, an increase or decrease of thephotoluminescent signal can be detected. By way of example, if thereaction product is H⁺ and the reactive moiety, such as imidazole, iscapable of binding the H⁺ to form the conjugate acid, the permeabilityof the ligand shell to a 9,10-anthraquinone-2-sulfonate quencher willincrease as the equilibrium concentration for the protonated moietyincreases. This in turn will affect the detectable photoluminescentsignal as the probability of electron transfer between the quencher andthe photoluminescence core increases with increasing permeability.

In some cases, the dynamically evolving molecular environment involvesthe depletion of a reactant. In cases where the reactive ligand in theligand shell is capable of reacting with a reactant to modulate thepermeability of the ligand shell, the sensor is capable of indirectlydetecting the formation of a product. As the reaction progresses andproduct is being formed, reactants are being consumed. The lowerconcentration of reactants will change the reaction equilibrium betweenthe first and second states. Depending on how the permeability of theligand shell is modulated, an increase or decrease of thephotoluminescent signal can be detected. By way of example, if thereactant is H⁺ and the reactive moiety, such as imidazole, is capable ofreversibly binding the H⁺ to form the conjugate acid, the permeabilityof the ligand shell to a 9,10-anthraquinone-2-sulfonate quencher willdecrease as the equilibrium concentration for the deprotonated moietyincreases. This in turn will affect the detectable photoluminescentsignal as the probability of electron transfer between the quencher andthe photoluminescence core decreases with decreasing permeability.

In some embodiments, the reactant or product may comprise H⁺, OH⁻, anacid, or a base. In some embodiments, the reactant may comprise anacetyl, a methyl, a phosphoryl, carboxyl, hydroxyl, or amine moiety.When the sensor comprises a reactive moiety capable of reacting with anyof H⁺, OH⁻, an acid, a base, an acetyl, a methyl, a phosphoryl,carboxyl, hydroxyl, or amine, the reaction between the

Miscellaneous

Unless otherwise specified or indicated by context, the terms “a”, “an”,and “the” mean “one or more.” For example, “a molecule” should beinterpreted to mean “one or more molecules.”

As used herein, “about”, “approximately,” “substantially,” and“significantly” will be understood by persons of ordinary skill in theart and will vary to some extent on the context in which they are used.If there are uses of the term which are not clear to persons of ordinaryskill in the art given the context in which it is used, “about” and“approximately” will mean plus or minus ≤10% of the particular term and“substantially” and “significantly” will mean plus or minus >10% of theparticular term.

As used herein, the terms “include” and “including” have the samemeaning as the terms “comprise” and “comprising.” The terms “comprise”and “comprising” should be interpreted as being “open” transitionalterms that permit the inclusion of additional components further tothose components recited in the claims. The terms “consist” and“consisting of” should be interpreted as being “closed” transitionalterms that do not permit the inclusion additional components other thanthe components recited in the claims. The term “consisting essentiallyof” should be interpreted to be partially closed and allowing theinclusion only of additional components that do not fundamentally alterthe nature of the claimed subject matter.

All methods described herein can be performed in any suitable orderunless otherwise indicated herein or otherwise clearly contradicted bycontext. The use of any and all examples, or exemplary language (e.g.,“such as”) provided herein, is intended merely to better illuminate theinvention and does not pose a limitation on the scope of the inventionunless otherwise claimed. No language in the specification should beconstrued as indicating any non-claimed element as essential to thepractice of the invention.

All references, including publications, patent applications, andpatents, cited herein are hereby incorporated by reference to the sameextent as if each reference were individually and specifically indicatedto be incorporated by reference and were set forth in its entiretyherein. Preferred aspects of this invention are described herein,including the best mode known to the inventors for carrying out theinvention. Variations of those preferred aspects may become apparent tothose of ordinary skill in the art upon reading the foregoingdescription. The inventors expect a person having ordinary skill in theart to employ such variations as appropriate, and the inventors intendfor the invention to be practiced otherwise than as specificallydescribed herein. Accordingly, this invention includes all modificationsand equivalents of the subject matter recited in the claims appendedhereto as permitted by applicable law. Moreover, any combination of theabove-described elements in all possible variations thereof isencompassed by the invention unless otherwise indicated herein orotherwise clearly contradicted by context.

EXAMPLES

Non-Covalent Control of the Electrostatic Potential of Nanoparticlesthrough the Formation of Interfacial Ion Pairs

Herein we describe the control of the electrostatic potential at theinterface between a colloidal semiconductor quantum dot (QD) and thesolvent, water, by tuning non-covalent interactions between the ligandshell of the QD and screening counterions. The electrostatic potentialof a nanoparticle, governed by charges of the ionizable tail groups ofthe particle's passivating organic ligands and electrostatic screeningfrom surrounding electrolyte, controls (i) the particle's solubilitywithin polar media such as water, (ii) the particle's tendency toself-assemble into superlattices and functional materials, (iii) thewettability, permeability and binding affinity of the particle's surfacewith respect to charged small molecules or macromolecular species (andtherefore the ability of its passivation layer to act as a selectivemolecular recognition platform for charged analytes and potentialadsorbates), and (iv) the particle's interaction with biomembranes. Theconcentration and distribution of non-covalently bound counterionsaround a particle's surface additionally influence the apparentdissociation constant of ionizable ligands immobilized on the surface byscreening inter-ligand interactions.

The electrical double layer model, widely applied through bothanalytical and simulation approaches, has provided quantitativepredictions of how the size, valence and concentration of surroundingions contribute to the inversion and/or amplification of surface chargessurrounding a macroion like a nanoparticle. This model does not,however, adequately describe the electrostatic screening of such ionswhen that screening has contributions from molecular-level interactionssuch as van der Wools interactions, which depend on the specificchemical structure and the hydrophilicity/hydrophobicity of both thesurface-bound ligands and the electrolytes.

Quantum dots, in addition to their practical applications as biologicalprobes, are excellent model systems to investigate the role of suchcomplex screening effects because of the sensitive response of theirexciton dynamics, which we can readily map with steady-state andtime-resolved optical spectroscopy, to the surrounding dielectricenvironment or to proximate small molecules. Here, we demonstratecontrol over the local electrostatic environment ofnear-infrared-absorbing and -emitting PbS QDs through non-covalentinteractions of their 6-mercaptohexanoate (MHA) ligand shell withproximate tetraalkylammonium (NR₄ ⁺) cations, FIGS. 1A and 1B. Theseinteractions consist of electrostatic and van der Waals components, andregulate the permeability of the anionic ligand shell of the QDs to anegatively-charged small molecule, 9,10-anthraquinone-2-sulfonate (AQ,FIG. 1B), as measured by the yield of photoinduced electron transferfrom the QD to AQ. We show, through photoluminescence (PL) experiments,free energy scaling analysis, and molecular dynamics simulations that,even though the cations are in fast exchange between the fully solvatedstate and the ion pair state, they effectively screen the repulsiveinteractions between the MHA and AQ and increase the probability that AQwill permeate the ligand shell. The efficacy of this screening increasesas the length of the alkyl chains of the cation (R) increases, becauseaccumulative van der Waals interactions between the backbones of MHAligands and the alkyl chains of NR₄ ⁺ increase the probability of ionpairing at the ligand/water interface.

This demonstration of the use of non-covalent interactions to controlthe electrostatic potential of a QD is important because non-covalentchemistry imparts great tenability to the properties of QDs in aqueousdispersions without the need for additional covalent functionalizationof their surfaces. Furthermore, this work demonstrates the sensitivityof our photoinduced charge transfer method to quantitatively probe theproperties of the QD/solvent interface: here, that techniquesuccessfully measures even the very weak, transient interactions betweenQDs and counterions, a challenge for structural and chemicalcharacterization tools like NMR.

Synthesis of MHA-capped PbS QDs and Quantification of Their LigandShells. We synthesized oleate-capped PbS QDs with a first excitonic peakat 968 nm, which corresponds to a core radius of 1.6 nm. We preparedwater-soluble PbS QDs capped with an adlayer of MHA by adding 400 molarequivalents of MHA to displace the native oleate ligands andsubsequently transfer the QDs to water, a procedure detailed in ourprevious work [He, C.; Weinberg, D. J.; Nepomnyashchii, A. B.; Lian, S.;Weiss, E. A. J. Am. Chem. Soc. 2016, 138, 8847-54]. We used ¹H NMR toquantify the composition of the ligand shell of the MHA-capped QDs, and,based on five measurements on separately prepared samples, each PbS QDhas an average of 102±11 MHA ligands bound to its surface. No boundoleate was detected on the surfaces of the water-soluble QDs.

The ability of NR₄ ⁺ counterions to screen QD-molecule repulsionincreases with bulkier R. We prepared a series of 2.63 μM samples ofMHA-capped QDs mixed with 1000 equiv. of AQ and increasing molar equiv.(0-2.0×10⁵ per QD) of NR₄ ⁺, where R=—CH₃, tetramethylammonium (TMA);R=—CH₂CH₃, tetraethylammonium (TEA); R=—(CH₂)₂CH₃, tetrapropylarnmonium(TPA); or R=—(CH₂)₃CH₃, tetrabutylammonium (TBA). We allowed all thesamples to equilibrate in the dark for four hours before performing anyoptical measurements. AQ is an electron acceptor with respect tophotoexcited PbS QDs. The probability of electron transfer (eT) from thephotoexcited QD to a proximate AQ is a measure of the permeability ofthe MHA ligand shell to the AQ. We measure this probability directlyfrom the degree of photoluminescence (PL) quenching of the QD ensembleupon addition of AQ, since eT from the QD to AQ results in acharge-separated state that recombines non-radiatively. With no NR₄ ⁺present, the free energy of transfer of the charged AQ from the bulksolution to the inorganic surface of the QD increases by 154 J/mol uponintroduction of each additional charged ligand to the ligand shell.

In FIG. 2A, we plot “PL/PL₀” vs. the concentration of NR₄ ⁺ in the QDsolution, where “PL” is the integrated photoluminescence intensity ofeach sample containing 1000 equiv. of AQ and a given concentration ofNR₄ ⁺, and “PL₀” is the intensity of each sample with the same NR₄ ⁺concentration but with no added AQ. The ratio “PL/PL₀” is therefore thefraction of the emissive population of QDs that remain emissive afteraddition of the quencher AQ. For all the cation structures studied, theAQ quenches the PL of the QDs more effectively—that is, the yield of eTincreases—as the concentration of the screening counterion NR₄ ⁺increases; NR₄ ⁺ therefore increases the permeability of the ligandshell to AQ. The effectiveness of this screening depends on the size andstructure of the counterion: the maximum yield of eT, i.e., (1−theminimum PL/PL₀), increases from ˜50% to ˜80% as we increase the numberof carbons in the R groups of the cation from 1 to 4. The screening bythe counterion is therefore more effective as the steric bulk of thecounterion increases (while holding the charge constant).

Transient absorption (TA) experiments on QD-AQ mixtures allow us toidentify two general mechanisms of eT in this system, and to determinewhich is more sensitive to screening by NR₄ ⁺. Addition of 1000 equiv.of AQ to PbS QDs accelerates the decay of the QD exciton, monitored asthe recovery of the ground state bleach feature of the QD at 1014 nm, onboth the nanosecond timescale and the microsecond timescale when no NR₄⁺ is present. We assign the extracted single-ns decay process introducedby addition of the AQ to eT from the photoexcited QD to statically boundAQ molecules on its surface. We assign the increase in the rate ofexciton decay from (1 μs)⁻¹ to (0.7 μs)⁻¹ shown in FIG. 2B todiffusion-controlled eT from the QD to AQ.

The rate and fractional amplitude of the single-ns eT process is onlyslightly affected by the presence of NR₄ ⁺ counterions and showsnegligible dependence on the bulkiness of these species, For instance,upon addition of 50,000 equiv. of TMA to the QD-AQ mixture, this timeconstant is 1.4 ns, and upon addition of the same amount of TBA to theQD-AQ mixture, this time constant is 1.3 ns.

The eT from the QDs to AQ on the μs timescale is, in contrast, verysensitive to the presence and identity of the NR₄ ⁺ counterions. Whileaddition of TMA seems to have negligible effect on the rate of eT onthis timescale (compare black and red traces in FIG. 2B), addition of50,000 equiv. of TBA reduces the exciton lifetime of QDs within QD-AQmixtures by a factor of three, compare black and blue traces in FIG. 2B,and see Table 1. This result is consistent with the dramatic increase(from ˜35% to ˜80%) in the yield of eT from QDs to AQ upon addition ofTBA measured from PL quenching experiments. It appears then, that theaddition of NR₄ ⁺ predominantly affects the rate and yield ofdiffusion-controlled eT from the QD to AQ, rather than that of eT withinstatic QD-AQ complexes. This result is reasonable since, while staticQD-AQ complexes form from occupation of AQ in gaps within the ligandshell, the rates of diffusion-controlled eT processes depend on the rateof transport of the charged AQ across the ligand/solvent interface, andthis transport depends on the electrostatic interactions at thisinterface.

Importantly, the presence NR₄ ⁺ does not affect the excited statedynamics of QDs in the absence of AQ, so the acceleration of theirexciton decay in the presence of AQ can be attributed exclusively to theacceleration of the eT processes from QDs to AQ.

TABLE 1 Time Constants for Decay of the QD Exciton in the Presence of1000 equiv. AQ, on the Nano-to-Microsecond Timescale. Amplitude-weightedτ1 (ns)^(a) τ2 (μs)^(a) Average Time Constant, Sample (A1) (A2) τ(μs)^(b) PbS QD + AQ 96 ± 26 0.70 ± 0.03 0.59 (−0.19) (−0.81) PbS QD +AQ + 51 ± 12 0.61 ± 0.07 0.53 50,000 equiv. TMA (−0.15) (−0.85) PbS QD +AQ + 10 ± 1  0.35 ± 0.01 0.22 50,000 equiv. TBA (−0.39) (−0.61)^(a)Error bars are fitting errors from the kinetic traces extracted (at1014 nm) from the nanosecond-to-microsecond TA spectrum of each sample.^(b)Calculated.

Both steady-state PL and TA measurements indicate that the ability ofthe cation to screen the electrostatic repulsion between the QD'sligands and AQ increases with increasing size of the cation. Here,screening is more effective for bulkier counterions. The counterionshave some hydrophobic portion that can drive ion-pairing with thehydrophobic portion of the ion it screens through favorable van derWaals interactions.

Without wishing to be bound by theory, we believe that the trend weobserve—that TBA screens repulsion between QD and AQ more effectivelythan TMA—is due to these interactions. Several factors potentiallycontribute to the enhanced electrostatic screening effects observed withthe increasing size of NR₄ ⁺. As R increases in length, NR₄ ⁺ forms athicker condensation layer on the QD surface. The bigger condensedcations, TEA, TPA and TBA, penetrate the MHA ligand shell and formmultiple layers on the QD surface. Second, as the attraction strength,ϵ, between the —CH₂+ groups in NR₄ ⁺ and MHA increases, the number ofhydrophobic contacts, n_(c), between the methylene groups of MHA and NR₄⁺ increases for all the cations. Third, the location of the slippingplane (i.e., the location where the number density of NR₄ ⁺ cationsdecays to its bulk value) shifts away from the QD surface as the size ofNR₄ ⁺ increases and their condensation layer increases in thickness.Fourth, the integrated net charge per unit area, Q(z), at the slippingplane decreases in magnitude as the alkyl chains of NR₄ ⁺ increase inlength, which suggests that bulkier NR₄ ⁺ cations are more effective inscreening the electrostatic potential of a negatively charged QD. Fifth,density of the charged ligands on the surface may allow for the cationsto penetrate the ligand shell. Sixth, the length of the charged ligandmay affect the degree of ordering within the ligand corona.

In summary, the electrostatic potential at the surfaces of PbS QDs,passivated by an anionic ligand shell of 6-mercaptohexanoate, can bereadily adjusted by non-covalent interactions with non-coordinatingtetraalkylammonium cations (NR₄ ⁺) though the formation of interfacialion pairs. By changing the length of the alkyl chains on these cationsand adjusting their concentrations, we regulate the permeability of theanionic ligand shell of QDs to a negatively charged molecular redoxprobe, 9,10-anthraquinone-2-sulfonate. The ability of NR₄ ⁺ to screenthe repulsive interaction between the negatively charged QDs and themolecular probe increases as either the concentration or the stericbulkiness of NR₄ ⁺ increases. The probability of forming an interfacialion pair is regulated by not only the number of available counterions inbulk solution, but also the intrinsic binding affinity of eachligand-counterion pair. This binding affinity is influenced by bothelectrostatic and van der Waals contributions toward the total freeenergy of the system. As a result, non-covalent interactions expands thecurrent toolset of nanoparticle surface chemistry, and suggestsstrategies for implementing a combination of nanoscale forces, such aselectrostatic/van der Waals interactions and entropic effects, in thedevelopment of nanoparticle-based platforms in self-assembly, molecularrecognition, and photocatalysis.

Synthetic Procedures for Oleate-Capped PbS QDs. We synthesized 1.6 nmoleate-capped PbS QDs using a procedure adapted from that of Hines andScholes [Hines, M. A.; Scholes, G. D., Adv. Mater. 2003, 15, 1844-1849].We mixed 0.36 g PbO and 2.0 mL oleic acid (OA) with 18.0 mL 1-octadecene(ODE) in a 50-mL three-neck round bottom flask at room temperature.Heating the mixture up to 150° C. with constant stirring under N₂ flowfor an hour produced a clear and colorless solution. We cooled themixture to 110° C., and injected 0.17 mL of hexamethyldisilathianedissolved in 8 mL of ODE. The solution turned from orange to brownwithin 3 seconds. After 10 minutes, we used an ice bath to cool thereaction mixture to room temperature. The product was separated intofour 50-mL centrifuge tubes for further purification. We purified theQDs by first washing the reaction mixture with acetone (6:1 by volume),and centrifuging it at 3500 rpm for 20 min. We then decanted thesupernatant, dried the QD pellet, redispersed the QDs in 7.5 mL hexanes,and precipitated the QDs two additional times, as described above, using12.5 mL methanol and acetone, respectively, as the non-solvents. Thecleaned PbS QDs were finally dispersed in a minimal amount of hexanes toform the stock solution.

Synthesis of MHA-capped PbS QDs and Quantification of Their LigandShells. We synthesized oleate-capped PbS QDs with a first excitonic peakat 968 nm, which corresponds to a core radius of 1.6 nm based on theempirical formula reported by Moreels et al., ACS Nano 2009, 3, 3023-30,using a procedure adapted from that of Hines and Scholes, Adv. Mater.2003, 15, 1844-1849, We prepared water-soluble PbS QDs capped with anadlayer of MHA by adding 400 molar equivalents of MHA to displace thenative oleate ligands and subsequently transfer the QDs to water, aprocedure detailed in our previous work³⁴ and in the SupportingInformation. We used ¹H NMR to quantify the composition of the ligandshell of the MHA-capped QDs, and, based on five measurements onseparately prepared samples, each PbS QD has an average of 102 ±11 MHAligands bound to its surface. No bound oleate was detected on thesurfaces of the water-soluble QDs.

Sizing of PbS QDs via Ground State Absorption and Transmission ElectronMicroscopy. The ground state absorption spectrum of a 4.1 μM solution ofoleate-capped PbS QDs was obtained on a Varian Cary 5000 spectrometerusing a 2 mm/10 mm dual pathlength quartz cuvette. We corrected thebaseline of the spectrum with hexanes prior to measurement, anddetermined the size of the synthesized PbS QDs (and their respectiveextinction coefficient) from the position of the first excitonic peak(˜968 nm) using the calibration curve published by Moreels et al., ACSNano 2009, 3, 3023-30. All concentrations of QDs were calculated fromthe absorbance of QDs at 400 nm.

In order to verify the accuracy of this calibration technique, weperformed transmission electron microscopy experiments using a JEOLJEM-2100F FAST TEM. We prepared TEM samples by drop-casting a solutionof PbS QDs in hexanes onto a Carbon Type B film (Ted Pella, Inc.). Weanalyzed 81 PbS QDs using the ImageJ software package, and determinedthat the average diameter of these particles is 2.9±0.3 nm. The 1.6 nmradius for oleate-capped QDs that we calculated from absorptionspectroscopy is within the error of our TEM measurements.

Preparation of 6-mercaptohexanoate (MHA)-capped PbS QDs by LigandExchange. We prepared water-soluble PbS QDs capped with MHA throughligand exchange. [He, C. et al., J. Am. Chem. Soc. 2016, 138, 8847-54].We added 400 equiv. of MHA to a 5 mL sample of 40 μM oleate-capped PbSQDs dispersed in CHCl₃, and shook the mixture rigorously for 1 min untilthe QDs flocculated. We then added 480 equivalents of NaOH per QD(NaOH/MHA=1.2:1) to the mixture to deprotonate the —COOH groups(pK_(a)≈4.8), and make the QDs negatively charged and water-soluble. TheQDs precipitated out of solution as we added NaOH, and transferred tothe aqueous layer as we added 4 mL of water on top of the chloroform andgently shook the mixture. We then centrifuged this mixture at 7000 rpmfor 10 min to facilitate the separation between aqueous and organiclayers, which are sometimes emulsified due to the presence ofsurfactants. The optically clear aqueous layer was separated and washedwith 10 mL chloroform to eliminate displaced oleate species, and thisaqueous layer served as a stock solution of MHA-capped PbS QDs.

Quantification of MHA Ligands within the Ligand Shell of PbS QDs. Weprepared MHA-capped, water-soluble PbS QDs using the proceduresdescribed above, determined their concentration from the intensity oftheir ground state absorption spectra at 400 nm, and applied ¹NMR toquantify the number of bound MHA ligands per QD. The NMR samples were39.5 μM solutions of the QDs with 800 equiv. of sodium formate as aninternal integration standard (singlet at 8.36 ppm, 1H). We set theacquisition time to 30 s and the relaxation time to 90 s, respectively,to allow for complete collection of the free induction decay signal andsufficient relaxation of proton nuclei between measurements, andperformed 32 scans to get a spectrum with satisfactory signal-to-noiseratio. The 2.01-2.28 ppm region of the resulting spectra (which containssignal from the methylene protons alpha to the carboxylate group in MHA)is fit with a sum of Lorentzian functions, and the broad featurecentered at ˜2.13 ppm, which corresponds to those protons of MHA ligandsthat are bound to the surface of QDs, is integrated against the sodiumformate internal standard. The quantitative NMR analyses are tabulatedin Table 2.

TABLE 2 Compositions of the Ligand Shells of MHA-Capped PbS QDs. No. ofSample No. of Bound MHA per QD 1 95 2 90 3 112 4 97 5 117Average/Standard Deviation 102 ± 11

Addition of NR₄ ⁺ chloride salts has negligible impact on the electronicstructure and colloidal stability of MHA-capped PbS QDs. Ground-stateabsorption spectra collected from a 2.63 μM aqueous solution ofMHA-capped PbS QDs, mixed with 200,000 equiv. of TMA, TEA, TPA or TBAwere superimposed. The backgrounds of these spectra were subtractedusing the absorbance of an aqueous solution of the respective NR₄ ⁺chloride salt with the same concentration. The four spectra overlaysatisfactorily with each other with no sign of scattering in eithervisible or the near infrared region, which indicates that the additionof NR₄ ⁺ chloride salts has negligible effects on the electronicstructure and colloidal stability of MHA-capped PbS QDs.

Representative Photoluminescence Spectra of MHA-Capped QDs upon Mixingwith Increasing Molar Equivalents of NR₄ ⁺ Chloride Salts. We preparedthe samples for PL measurements using the procedures described above.This series of samples have no AQ added and serve as the blank controls.All the samples were contained in a 10 mm/2 mm dual path length quartzcuvette and photoexcited with an 800-nm beam along the 10-mm axis, andthe corresponding PL spectra were collected in a right-angle geometry,FIG. 8. The integrated PL intensity, which we define as “PL₀” in themain text, FIG. 2A, is calculated from integrating each spectrum afterbaseline subtraction. In some cases, the PL intensity of PbS QDs isslightly enhanced upon addition of NR₄ ⁺ chloride salts. Thisenhancement in PL quantum yield potentially results from the passivationof surface electron trap states by Cl⁻ anions.

Representative Photoluminescence Spectra of MHA-Capped QDs upon Mixingwith Increasing Molar Equivalents of NR₄ ⁺ Chloride Salts and 1000equiv. AQ. We recorded the PL spectra of these samples using the samesetup described earlier. The integrated PL intensity, which we define as“PL”, is calculated from integrating each spectrum after baselinesubtraction, and normalized to the integrated PL intensity of theirrespective blank samples, “PL₀”, to divide out the minor fluctuations inPL intensity caused by NR₄ ⁺ chlorides alone.

Diffusion-Ordered Spectroscopy (DOSY) NMR determines the hydrodynamicradii of NR₄ ⁺ cations. We recorded DOSY spectra of NR₄ ⁺ chloride salts(TMA, TEA, TPA and TBA, 0.1 M) in D₂O using a double stimulated echoexperiment with bipolar gradients (“dstebpgp3s” sequence) on a BalkerAvance-III 600 MHz NMR spectrometer. We used a diffusion delay, Δ, of0.2 s, a gradient length, δ, of 2000 μs, and chose 16 gradient strengthvalues from 5% to 60% for all our measurements. In order to determinethe diffusion coefficients and hydrodynamic radii of these species, weplotted the dependence of the integrated ¹H signal of themethylene/methyl protons alpha to N (3.10 ppm) as a function of thegradient function, G², given by eq 1,

$\begin{matrix}{G^{2} = {\left( {\gamma \; g\; \delta} \right)^{2}\left( {\Delta - \frac{\delta}{3}} \right)}} & (1)\end{matrix}$

where γ is the gyromagnetic ratio of a proton (2.68×10⁴ s⁻¹G⁻¹) and g isthe gradient strength (G/cm). We performed a fit of these curves usingeq 2,

I(G ²)=I ₀ e ^(−DG) ² =I ₀ e ^(−G) ² ^(/m)  (2)

to determine the diffusion coefficients, D, and calculated thehydrodynamic radii, r, using the Stokes-Einstein equation, eq 3,

$\begin{matrix}{r = {\frac{k_{B}T}{6{\pi\eta}\; D} = \frac{k_{B}{Tm}}{6{\pi\eta}}}} & (3)\end{matrix}$

where k_(B) is the Boltzmann constant (1.38×10⁻²³ J/K), T is thetemperature (298 K), η is the viscosity of D₂O (0.89 mPa·s), and m=1/D.The fitting results can be found in Table 3.

TABLE 3 Fitting Parameters for DOSY Experiments on TetraalkylammoniumCations. 1/Diffusion Diffusion Name of Coefficient CoefficientHydrodynamic species (m, s/cm²)^(a) (D, × 10⁻⁶ cm²/s) Radius (r, nm)^(b)TMA 96443 ± 1389 10.4 0.237 ± 0.003 TEA 130929 ± 390  7.64 0.321 ± 0.001TPA 184517 ± 520  5.42 0.453 ± 0.001 TBA 224341 ± 945  4.46 0.550 ±0.002 ^(a)Error bars are fitting errors. ^(b)Errors are propagateddirectly from the fitting errors in m using the equation${\Delta r} = {\frac{k_{B}T}{6{\pi\eta}} \times {{\Delta m}.}}$

NR₄ ⁺ cations are in fast exchange between the fully-solvated state andthe ion pair state when mixed with oppositely charged QDs. We preparedtwo samples where TBA chloride salt is either dissolved in pure D₂O ormixed with a 2.63 μM sample of MHA-capped PbS QDs (TBA:QD=50:1). Werecorded DOSY spectra on these samples using the procedures describedabove, and applied eqs 2 and 3 to extract the diffusion coefficient andhydrodynamic radius of TBA cations. We observe that both spectra arewell-fit by a single-exponential function, which indicates that there'sconsistently one single population of TBA cations in solution; ii) thehydrodynamic radius of TBA mixed with QDs shows an increase of up to 12%compared to the sample without any added QDs, see Table 4. We thereforeconclude that TBA cations are in fast exchange on and off the QDsurface, a process that cannot be deconvolved within the temporalresolution of NMR and contributes to a weighted-average of hydrodynamicradii between the two states.

TABLE 4 Fitting Parameters for DOSY Experiments on TBA Chloride inAqueous Solution. 1/Diffusion Diffusion Coefficient^(a) CoefficientHydrodynamic Name of sample (m, s/cm²) (D, × 10⁻⁶ cm²/s) Radius^(b) (r,nm) QD + 50 TBA 250843 ± 3338 3.99 0.615 ± 0.008 TBA only 224341 ± 945 4.46 0.550 ± 0.002 ^(a)Error bars are fitting errors. ^(b)Errors arepropagated directly from the fitting errors in m using the equation${\Delta r} = {\frac{k_{B}T}{6{\pi\eta}} \times {{\Delta m}.}}$

The ion-pairing effect between AQ and NR₄ ⁺ is not the major contributorto the increasing yield of PL quenching in FIG. 1A. We recorded DOSYspectra on five 2.63 mM samples of AQ mixed with increasing equiv.(0-200 per AQ) of TBA chloride salt in D₂O using the same pulse sequencedescribed above, and calculated the diffusion coefficient andhydrodynamic radius of AQ in each sample using eqs 2 and 3, see Table 5.Here we note that the concentration of AQ and the range ofconcentrations for TBA are the same as those samples we used for PLmeasurements. The hydrodynamic radii of AQ are plotted with PL/PL₀ (fromFIG. 2A), as a function of the TBA/AQ ratio, see FIG. 3. The lack ofresemblance between these two curves suggests that it is the ion-pairingeffect between the QD ligand shell and NR₄ ⁺, not the ion-pairing effectbetween AQ and NR₄ ⁺, that results in the enhanced yield of electrontransfer between MS QDs and AQ.

TABLE 5 Fitting Parameters for DOSY Experiments on a 2.63 mM Solution ofAQ Mixed with Increasing Equiv. of TBA Chloride. 1/Diffusion DiffusionCoefficient Coefficient Hydrodynamic Name of sample (m, s/cm²)^(a) (D, ×10⁻⁶ cm²/s) Radius (r, nm)^(b) AQ Only 165639 ± 3842 6.04 0.406 ± 0.009AQ + 5 TBA 190424 ± 2112 5.25 0.467 ± 0.005 AQ + 10 TBA 195676 ± 18175.11 0.480 ± 0.004 AQ + 50 TBA 258552 ± 4484 3.87 0.634 ± 0.011 AQ + 200TBA  454235 ± 27021 2.20  1.11 ± 0.066 ^(a)Error bars are fittingerrors. ^(b)Errors are propagated directly from the fitting errors in musing the equation${\Delta r} = {\frac{k_{B}T}{6{\pi\eta}} \times {{\Delta m}.}}$

Transient Absorption Measurements. Picosecond-to-nanosecond TransientAbsorption Spectroscopy. We split the 2.5-mJ output of a commercialamplified Ti-sapphire laser (Solstice, 1 kHz, 100 fs, Spectra Physics),and guided 95% to an optical parametric amplifier (TOPAS-C, LightConversion) used to produce the pump wavelength (850 nm) for sampleexcitation, and 5% to a commercial TA spectrometer (HELIOS, UltrafastSystems) for use as the probe for TA experiments with pump-probe delaytimes up to 3200 ps. Within the spectrometer, a single filamentbroadband continuum of probe wavelengths from 900 to 1400 nm wasgenerated in a 1.2-cm thick sapphire plate and then passed through along-wave pass filter to isolate near-infrared wavelengths above 850 nm.The probe light was then split into sample and reference beams. Wecombined the sample beam with the pump light at the sample, which wascontained in a 2-mm quartz cuvette. The transmitted probe signal wascollected into an optical fiber and dispersed onto an array detector.Dividing the signal from the sample beam by the signal from thereference beam allowed us to divide out any fluctuations in the probebeam intensity during the experiment. The output differential absorptionspectrum (ΔA) was obtained through active background subtraction of theground state spectrum by chopping the pump at 500 Hz. The pump light wasdepolarized to prevent unintentional photoselection so that measurementsreflect only population dynamics, and we adjusted the power to 100 μW toavoid any multi-exciton effects. The solution was stirred with amagnetic stir bar to minimize local heating.

Nanosecond-to-microsecond Transient Absorption Spectroscopy. We used acommercial spectrometer (EOS, Ultrafast Systems) to collect TA spectrafor pump-probe delay times from 0.5 ns-10 82 s. The excitation beam wasgenerated by the same method and followed the same beam path as theexcitation used for the picosecond-to-nanosecond TA experiment describedabove. The proprietary EOS light source generates a super-continuum(400-1700 nm) probe pulse by focusing a diode laser into a photoniccrystal fiber. The repetition rate of the probe pulse is 2 kHz, which istwice the repetition rate of the pump pulse, and it is triggered in syncwith the pump pulse. The pump-probe delay is electronically generatedand measured by an electronic timer-counter-analyzer (Pendulum). Afterthe probe pulse passes through an 850-nm long pass filter, it is splitby a beam-splitter into sample and reference beams. The sample beam wassent onto the sample, which was contained in a 2-mm quartz cuvette, andfocused to a spot size of ˜200 μm. The reference and sample beams arethen collected in optical fibers and dispersed onto array detectors. Inorder to achieve reasonable signal to noise, the incident power of pumplight was adjusted to 2 mW. The solution was stirred with a magneticstir bar to minimize local heating.

We performed transient absorption experiments, as described above, on aset of separately prepared 6.58 μM PbS QDs. The compositions of thesesamples are described in the main text. These samples were allowed toequilibrate in the dark overnight before we took measurements. Weapplied a sum of exponentials convoluted with an instrument responsefunction (in this case, a Gaussian pulse) to fit the kinetic traces ofthe ground state bleach (extracted at 1014 nm) and determine therelaxation dynamics of photo-excited carriers on both ultrafast andnanosecond-to-microsecond timescales, eq 4.

$\begin{matrix}{y = {{IRF} \otimes {\sum\limits_{1}^{n}{A_{n}e^{\frac{- {({t - t_{0}})}}{\tau_{n}}}}}}} & (4)\end{matrix}$

Here n is the minimal number of exponential functions required toadequately fit the data, that is, to yield a symmetric, randomdistribution of residuals around the zero line.

For QD samples with no added AQ, the kinetic traces of the ground statebleach on a picosecond-to-nanosecond time scale are fit with a sum ofthree exponentials. These three time constants are fixed when we fit thekinetics from the same QD samples mixed with AQ (while their amplitudesare allowed to vary), and a fourth exponential is added to account forthe accelerated decay of QD excitons induced by electron transfer. Thefitting parameters are summarized in Table 6.

The nanosecond-to-microsecond kinetic traces of the ground state bleachfor QDs mixed with NR₄ ⁺ counterions and AQ are fit with a sum of twoexponential functions. We also fit the kinetic traces extracted from theTA spectra of QD samples with no added AQ, and the fitting parametersare tabulated in Table 6. These data show that the presence of NR₄ ⁺counterions alone has negligible effects on the exciton dynamics of QDs,and the excited state life time of these QDs is consistently ˜1 μs.

TABLE 6 Time Constants for Decay of the QD Exciton on aPico-to-nanosecond Time Scale. τ₁ (ps)^(a,b) τ₂ (ps)^(a,b) τ₃ (ps)^(a,c)τ₄ = τ_(eT) (ns)^(a,d) Sample (A₁) (A₂) (A₃) (A₄) PbS QD 2.5 ± 0.7 38.3± 8.9  >3000 — (−0.07) (−0.06) (−0.87) PbS QD + AQ 2.5 ± 0.7 38.3 ±8.9  >3000 2.2 ± 0.3 (−0.03) (−0.05) (−0.66) (0.23) PbS QD + TMA 3.4 ±1.3 31.3 ± 10.3 >3000 — (−0.05) (−0.04) (−0.90) PbS QD + AQ + 3.4 ± 1.331.3 ± 10.3 >3000 1.4 ± 0.1 TMA (−0.03) (−0.06) (−0.67) (0.23) PbS QD +TBA 3.4 ± 0.9 50.0 ± 14.0 >3000 — (−0.06) (−0.05) (−0.88) PbS QD + AQ +3.4 ± 0.9 50.0 ± 14.0 >3000 1.3 ± 0.1 TBA (−0.04) (−0.07) (−0.65) (0.22)^(a)Error bars are fitting errors. ^(b)These small-amplitude componentscorrespond to surface-mediated charge trapping processes. ^(c)Thiscomponent is well beyond the measurable time window (3 ns) of our TA setup and cannot be fit accurately, but is required to make the fittingsconverge. ^(d)This component corresponds to the electron exchangeprocess (marked by eT) between photoexcited QDs and AQ, the rate andmagnitude of which is not sensitive to the presence and structure of NR₄⁺ counterions. Here we note that this component could containcontributions from both the charge separation (electron transfer fromthe conduction band of QD to the LUMO of AQ) and the chargerecombination (recombination between the electron on AQ and the hole inthe valence band of QD) processes, but we are not able to deconvolvethem.

Calculations of Amplitude-Weighted Average Time Constants. Theamplitude-weighted average time constants, τ, for decay of the QDexciton on a nano-to-microsecond time scale are calculated using thefollowing equation:

$\begin{matrix}{\tau = \frac{\Sigma \mspace{14mu} A_{i}\tau_{i}}{\Sigma \mspace{14mu} A_{i}}} & (5)\end{matrix}$

where A_(i) and T_(i) are the amplitude (%) and individual time constantfor each component of excitonic decay (see Table 1 and Table 7), whichwe obtained from fitting the kinetic traces of TA spectra with a sum ofexponential functions.

TABLE 7 Time Constants for Decay of the QD Exciton on aNano-to-microsecond Time Scale. Amplitude-weighted τ₁ (ns)^(a,b) τ₂(μs)^(a) Average Time Constant, Sample (A₁) (A₂) τ^(c) (μs) PbS QD 184 ±35 1.14 ± 0.03 1.01 (−0.15) (−0.86) PbS QD + TMA 158 ± 52 1.06 ± 0.020.93 (−0.14) (−0.86) PbS QD + TBA  85 ± 18 1.03 ± 0.02 0.94 (−0.10)(−0.90) ^(a)Error bars are fitting errors. ^(b)Correspond tosurface-mediated charge trapping processes. ^(c)Calculated.

Control of Redox Activity by Tuning the Electrostactic Interactions atthe Nanoparticle-Solvent Interface

Herein we report the relationship between a controlled electrostaticcharge density within the organic adlayer of a colloidal PbS quantum dot(QD) and the permeability of that adlayer to a negatively charged smallmolecule, 9,10-anthraquinone-2-sulfonic add sodium salt (AQ). We coatthe PbS QDs with mixed monolayers of neutral ligands (6-mercaptohexanol,MHO) and ligands with a negatively charged tail group(6-mercaptohexanoate, MHA), and change the ratio of MHO to MHA withinthe monolayer to tune the Coulomb repulsion between the outer surface ofthe ligand shell and the charged sulfonate substituent on AQ. Thesenanoscale electrostatic interactions control the permeability of theligand shell to AQ, and the probability of photoinduced electrontransfer (eT) from the QD to AQ.

Measurement of the yield of photoinduced eT between the QD and AQ is asensitive probe of adlayer permeability. The yield of eT upon mixing,for example, 200 molar equivalents of AQ with the QDs ranges from 10% ata charge density of 1.1 charges/nm² to 98% at a charge density of 0.29charges/nm². The basis of this technique is that eT between a QD and asmall molecule acceptor does not occur on a timescale competitive withother relaxation mechanisms of the QD unless the small molecule haspermeated through the ligand shell and is at (or very near) theinorganic surface of the particle. The probability of eT is thereforedirectly correlated with the probability of permeation. The use ofphotoinduced interfacial eT, rather than cyclic voltammetry (which iscommonly used to probe the structure of monolayers on planar metal andmetal nanoparticles) for this measurement is necessary because appliedstatic potentials typically induce irreversible redox processes on QDsurfaces, whereas eT from the photoexcited state of the QD is reversibleand non-destructive.

This work allows us to rationally design monolayers of photoactive,electroactive, semiconductor QDs for nanoscale molecular recognitionbased on electrostatic interactions.

Synthesis of Water-Soluble PbS QDs. Oleate-capped PbS QDs with a firstexcitonic peak at ˜985 nm and radius of 1.6 nm were synthesized using aprocedure adapted from that of Hines and Scholes, Adv. Mater. 2003, 15,1844-1849. We prepared water-soluble PbS QDs capped with mixedmonolayers of 6-mercaptohexanoate (MHA) and 6-mercaptohexanol (MHO)through ligand exchange using a method adapted from those of Hyun etal., J. Phys. Chem. B 2007, 111, 5726-5730 and Kalsin et al., J. Am.Chem. Soc. 2007, 129, 6664-6665. We added 400 equivalents of thiols perQD in total, with various MHO/MHA ratios, to a 5 mL sample of 40 μMoleate-capped PbS QDs dispersed in CHCl₃, and shook the mixturerigorously for 1 min until the QDs flocculated. We then added between 96and 480 equivalents of NaOH per QD (NaOH/MHA=1.2:1) to the mixture todeprotonate the —COOH groups (pK_(a)≈4.8), and make the QDs negativelycharged and water-soluble. The QDs precipitated out of solution as weadded NaOH, and transferred to the aqueous layer as we added 4 mL ofwater on top of the chloroform and gently shook the mixture. We thencentrifuged this mixture at 7000 rpm for 10 min to facilitate theseparation between aqueous and organic layers, which are sometimesemulsified due to the presence of surfactants. The optically clearaqueous layer was separated and washed with 10 mL chloroform toeliminate displaced oleate species, and this aqueous layer served as astock solution of MHA/MHO-capped PbS QDs. The range of pH for all QDsamples (2.63 μM) we investigated was 9.6-10.3. ¹H NMR spectra of theaqueous QD dispersions show that all the oleate ligands that wereinitially bound to the QDs are displaced upon addition of 400equivalents of thiols.

Quantification of MHA/MHO Mixed-monolayer Ligand Shell. We preparedwater-soluble PbS QDs capped with mixed monolayers of MHO/MHA of sixdifferent compositions using the procedures described above, anddetermined their concentrations from the intensity of their ground stateabsorption spectra at 400 nm. We then prepared a 13.2 μM sample of eachtype of QD in D₂O and quantified the compositions of the ligand shellsof the QDs in each sample by ¹H NMR spectroscopy, with 800 equivalentsof sodium formate added as the internal integration standard (sharpsinglet at 8.33 ppm, 1H). We set the acquisition time to 27 s and therelaxation time to be 90 s to allow for complete collection of FIDsignal and sufficient relaxation of ¹H nuclei between measurements,performed 8 scans of each sample (except for samples 5 and 6, for whichwe took 32 scans in order to improve the S/N ratio), and used a sum of 5Lorentzian functions to fit the acquired spectra.

The NMR spectra of the MHO/MHA-capped QDs contain a broad peak centeredat ˜2.07 ppm corresponding to the protons alpha to the —COO⁻ in boundMHA and a sharp triplet centered at ˜2.03 ppm corresponding to thoseprotons on freely diffusing MHA. A weak singlet at ˜2.08 ppm is assignedto an unknown impurity; the intensity of this peak does not scale withthe concentration of MHA, and we subtract this feature from the spectrumbefore integrating any peaks. The total number of MHA (bound plus free)per QD in solution is calculated by integrating the portion of thespectrum containing both peaks. In order to deconvolute the bound andfree MHA signals, we integrate the broad feature and sharp triplet from1.96 to 2.20 ppm separately against the sodium formate internalstandard, and assign these two numbers as the number of bound and freeMHA per QD, respectively. Table 8 lists the results of our quantitativeNMR analysis for all six samples of MHA/MHO-capped PbS QDs withdifferent mixed monolayer compositions. We note that the total number ofMHA ligands per PbS QD, as measured by NMR, is in a few cases slightly(up to 9.6%) larger than the total equivalents of MHA we added, adiscrepancy that can be accounted for by (i) the systematic error of ourNMR measurement (˜+8%), and (ii) incomplete phase transfer: up to 15% ofoleate-capped QDs, either unreacted or only partially exchanged by MHA,are not present in the final aqueous solution, which increases theMHA:QD ratio.

TABLE 8 Compositions of the Mixed Organic Adlayers of MHA/MHO-Capped PbSQDs. Eq. of Eq. of MHA Eq. Bound Eq. Free Eq. Bound MHA/MHO (Bound + MHAMHA MHO No. of Added to Free) Measured Mea- Esti- sample PbS QDsMeasured^(a,b) (x)^(a,c) sured^(a,d) mated^(a,e) 1 400/0  438 ± 36 115 ±9  323 ± 27  0 2 320/80  311 ± 26 100 ± 8  210 ± 17 80 3 280/120 286 ±24 94 ± 8 192 ± 16 116 ± 8 4 240/160 263 ± 22 81 ± 7 182 ± 15 139 ± 7 5160/240 170 ± 14 67 ± 6 104 ± 9  153 ± 6 6  80/320 86 ± 7 31 ± 3 55 ± 5189 ± 3 ^(a)The errors are propagated from systematic error in the NMRmeasurement using the calibration plot. ^(b)Calculated from the sum ofthe bound (~2.07 ppm, broad feature) and free (~2.03 ppm, sharp triplet)MHA signals. ^(c)Calculated from the broad feature centered at ~2.07ppm. ^(d)Calculated from the triplet centered at ~2.03 ppm.^(e)Estimated as (220-no. of bound MHA). If fewer than (220-no. of boundMHA) MHO ligands were added, eq. of MHO bound = eq. of MHO added.

The number of MHO ligands bound per QD is more difficult to determinefrom their NMR spectra, because the MHO molecules are in fast exchangewith the QD surface, and therefore present a single broad feature at˜3.5 ppm. We estimate the number of bound MHO ligands per QD bysubtracting the number of bound MHA ligands per QD from 220, the totalnumber of thiol binding sites per QD, which we measured by titrating theoleate-capped QDs with hexanethiol and counting the number of displacedoleates. This method of ligand counting is indirect, but only the numberof bound MHA per QD (and not the number of bound MHO per QD) determinesthe charge density at the QD surface, so the absolute number of MHO perQD is not critical to the analysis that follows. There are no detectableoleate ligands bound to the surfaces of the QDs after phase transfer.

Photoinduced Electron Transfer Occurs from PbS QDs to Adsorbed AQs. Weadded a series of equivalents of AQ (20 eq.-6000 eq.), which has onenegative charge on its sulfonate group (pK_(a)≈−1.8³³), to 2.63 μM ofPbS QDs, and allowed all of the samples to sit in the dark for fourhours to equilibrate. The PL of the QDs decreased monotonically withincreasing equivalents of AQ added, FIG. 4A. Based on theelectrochemical potentials of AQ and the measured conduction and valenceband-edges of PbS QDs of this size, measured using photoemissionspectroscopy, electron transfer from the conduction band-edge (or LUMO)of the QD to the LUMO of AQ is the most likely mechanism for quenchingof the QDs' PL. Electron transfer has a driving force of −0.3 eV, butboth hole transfer and energy transfer from the QD to AQ areenergetically uphill and do not occur.

Transient absorption (TA) measurements on the QD/AQ mixtures confirmthat the PL quenching upon addition of AQ is due to photoinducedelectron transfer from the QD to AQ. We performed TA on thepicosecond-to-nanosecond timescale on two samples of QDs with differentsurface compositions, one with QDs capped with 115 MHA/QD (sample 1) andone with QDs capped with 31 MHA/QD (sample 6), each mixed with twodifferent concentrations of AQ, as shown in FIGS. 5A and 5B. We alsoperformed the same measurements for these two QD samples with no addedAQ. The inset of FIG. 5A shows a representative TA spectrum of 115MHA-capped PbS QDs, at 2 ps after excitation with a 850-nm pump pulse.The large negative feature is the ground state bleach, and we monitorits evolution in time to measure the dynamics of exciton decay in theabsence and presence of AQ. FIGS. 5A and 5B show these dynamics on thepicosecond-to-nanosecond timescale for the QDs with different surfacecompositions. Addition of AQ to the QDs induces additional decaypathways on the 100-ps timescale, and this new decay gets faster (andresults in a larger bleach recovery) as the number of equivalents of AQincreases. These observations are consistent with the PL quenching data(FIG. 4A), and are characteristic of depopulation of the excited stateby electron transfer (eT) from the exciton of the QD to the LUMO ofstatically adsorbed AQ, as we and others have seen for many QD-moleculecombinations. The observed rate constant for eT scales linearly with thenumber of adsorbed molecular acceptors. The intrinsic eT rate withineach QD-AQ pair was calculated to be 2.4±0.2×10⁹ s⁻¹ from fitting a plotof the observed eT rate vs. the number of bound AQs, λ, with a line.

The Yield of Photoinduced Electron Transfer is a Probe of thePermeability of the Charged Organic Adlayer to AQ. FIG. 4B containsplots of the ratio PL/PL₀—the fraction of emissive QDs in a sample thatremain emissive after addition of AQ—vs. the concentration of free AQ inthe sample, for QDs with the six surface compositions that we studiedwith NMR. For a given concentration of added AQ, PL/PL₀ increases withincreasing coverage of the charged MHA ligand; therefore, the yield ofeT to the charged AQ molecule (which is inversely proportional toPL/PL₀) decreases with increasing negative charge density on the QDsurface. This conclusion is supported by the data in FIGS. 5A, 5B, whichshow that exciton decay is accelerated much more dramatically in the QDswith fewer surface charges, despite the fact that more highly chargedQDs were mixed with a factor of 20 more AQ than the less-charged QDs.This decrease is not due to a change in the energetics of the chargetransfer reaction (changing the density of charges that are more than 1nm from the QD surface has a negligible effect on the energy of thephotoexcited electron), but rather reflects a decrease in the averagenumber of AQ molecules adsorbed to each QD surface, i.e., the number ofdonor-acceptor complexes formed. The probability of forming a QD-AQdonor-acceptor complex is related to the permeability of the MHA/MHOligand shell to AQ.

In summary, we fabricated a series of water-soluble PbS QDs capped bymixed monolayers of charged mercaptohexanoate and neutralmercaptohexanol, with a controllable interfacial charge density, anddemonstrated the differential permeability of these monolayers to acharged small-molecule, a sulfonate-functionalized anthraquinone, usingphotoinduced electron transfer between the QD and the anthraquinone as aprobe of their interaction. Our results demonstrate the sensitivity ofphotoinduced charge transfer as a probe of not only the local chemicalenvironment of a colloid, but also the intermolecular structure of itssurfactant layer, which is difficult to characterize using traditionalanalytical tools. By embedding electrostatic interactions withinQD-molecule assemblies, we could construct a highly selectiverecognition/reaction platform for ionic species based on their charges,and the use of water as the medium for this system makes these resultsan exciting step towards potential applications of water-soluble QDs inbiological imaging, environmental sensing and photocatalysts.

Synthetic Procedures for Oleate-capped PbS QDs. We synthesized 1.6 nmoleate-capped PbS QDs using a procedure adapted from that of Hines, M.A.; Scholes, G. D., Adv. Mater. 2003, 15, 1844-1849. We mixed 0.36 g PbOand 2.0 mL oleic acid (OA) with 18.0 mL 1-octadecene (ODE) in a 50-mLthree-neck round bottom flask at room temperature. Heating the mixtureup to 150° C. with constant stirring under N₂ flow for an hour produceda clear and colorless solution. We cooled the mixture to 110° C., andinjected 0.17 mL of hexamethyldisilathiane dissolved in 8 mL of ODE. Thesolution turned from orange to brown within 3 seconds. After 10 minutes,we used an ice bath to cool the reaction mixture to room temperature.The product was separated into four 50-mL centrifuge tubes for furtherpurification, as is described in the main text. We purified the QDs byfirst washing the reaction mixture with acetone (6:1 by volume), andcentrifuging it at 3500 rpm for 20 min. We then decanted thesupernatant, dried the QD pellet, redispersed the QDs in 5 mL hexanes,and precipitated the QDs two additional times, as described above, using12.5 mL methanol and acetone, respectively, as the non-solvents. Thecleaned PbS QDs were finally dispersed in a minimal amount of hexanes toform the stock solution.

Sizing of PbS QDs via UV-vis Ground State Absorption and TransmissionElectron Microscopy. All ground state absorption spectra of solutions ofoleate-capped PbS QDs were obtained on a Varian Cary 5000 spectrometerusing a 2 mm/10 mm dual pathlength quartz cuvette (in which we excitealong the 2 mm axis). We corrected the baselines of all spectra withhexanes prior to measurement, and determined the size of the synthesizedPbS QDs (and their respective extinction coefficient) from the positionof the first excitonic peak (˜985 nm) using the calibration curvepublished by Moreels, I. et al., ACS Nano 2009, 3, 3023-3030. Allconcentrations of QDs were calculated from the absorbance of QDs at 400nm. In order to verify the accuracy of this technique, we performedtransmission electron microscopy experiments using a JEOL JEM-2100F FASTTEM. We prepared TEM samples by drop-casting a ˜12 μM solution of PbSQDs in hexanes onto a Carbon Type B film (Ted Pella, Inc). We analyzed101 PbS QDs using the ImageJ³ software package, and determined that theaverage diameter of these particles is 3.6±0.4 nm. The 1.6 nm radius foroleate-capped QDs that we predicted from UV-Vis spectroscopy is withinthe error of our TEM measurement.

Cyclic Voltammetric (CV) and Ground State Absorption Study of9,10-Anthraquinone-2-Sulfonic Acid Sodium Salt (AQ). In order todetermine the redox potential of AQ, we first performed cyclicvoltammetry experiments using a three-electrode cell with a 3-mm radiusglassy carbon electrode as the working electrode, saturated Ag/AgCl asthe reference electrode, and platinum wire as the counter electrode.Measurements were taken with Princeton applied research potentiostatsusing 0.5 mM deaerated solutions of AQ, with 0.25 M NaNO₃ added as thesupporting electrolyte. The pH of the solution is adjusted to 10.0 usingNaOH to mimic the actual condition for PL quenching experiments(pH=9.6√10.3).

Cyclic voltammogram shows quasi-reversible reduction of AQ under variousscanning rates (0.05 V/s, 0.1 V/s, 0.25 V/s and 0.5 V/s). The observedreduction potential is −0.59 V vs. Ag/AgCl reference electrode, which,after correction using eq 6, corresponds to ˜−4.1 eV for the LUMO energyof AQ relative to vacuum.

$\begin{matrix}\begin{matrix}{{E\left( {{vs}.\mspace{14mu} {vacuum}} \right)} = {{- {E\left( {{vs}.\mspace{14mu} {NHE}} \right)}} - {4.43\mspace{14mu} {eV}}}} \\{= {{- \left\lbrack {{E\left( {{{vs}.\mspace{14mu} {saturated}}\mspace{14mu} {Ag}\text{/}{AgCl}} \right)} + {0.199\mspace{14mu} {eV}}} \right\rbrack} - {4.43\mspace{14mu} {eV}}}} \\{= {{- \left\lbrack {E_{{reduction}\mspace{14mu} {peak}\mspace{14mu} {potential}} + {0.0285\mspace{14mu} {eV}} + {0.199\mspace{14mu} {eV}}} \right\rbrack} - {4.43\mspace{14mu} {eV}}}}\end{matrix} & (6)\end{matrix}$

UV-Vis ground state absorption spectrum of AQ (collected at aconcentration of 260 μM) shows that the optical transition with theminimal energy is centered at ˜330 nm, which translates into a 3.8 eVexcitation energy between the HOMO and the LUMO. Therefore, we subtractthis energy from the LUMO energy that we measured using CV, anddetermine that the HOMO energy of AQ is ˜−7.9 eV.

Ground state absorption spectra show that up to 15% of PbS QDs are lostduring the preparation of MHA/MHO-capped PbS QDs. We used UV-Visspectrophotometer to determine the concentration of PbS QDs by theamplitude of their ground state absorbance at 400 nm. Then we comparedthe total amount of PbS QDs present in solution before and after ligandexchange, and calculated the percentage of PbS QDs lost to be between14.4% and 0.0% for samples 1-6.

All of the native oleate ligands are displaced upon adding 400 eq. ofthiols to PbS QDs during the ligand exchange. FIG. 6A shows the fullspectra of MHA/MHO-capped water-soluble PbS QDs (13.2 μM), with sixdifferent compositions in their ligand shells. FIG. 6B compares therepresentative NMR spectrum of water-soluble QDs (after exchange) witholeate-capped QDs (before ligand exchange). For QDs that are dispersedin aqueous solution, the absence of signal from bound oleate (broadfeature centered at 5.69 ppm) indicates that all the native oleateligands are completely displaced upon adding 400 eq. of thiols.

Quantification of MHA and MHO within the Mixed Monolayer Ligand Shell ofWater-soluble PbS QDs. The ¹H NMR spectrum shows the signals from vinylprotons of bound oleate (5.69 ppm, broad) and free oleate species (5.50ppm, sharp multiplet) of oleate-coated QDs treated with differentconcentrations of hexanethiol (HT). As the amount of added HT increases,the signal from the vinyl protons of bound oleate decreases, and thesignal from the vinyl protons for freely-diffusing oleate speciesincreases. The sum of the integrated signals corresponding to bound andfree populations of oleate species (measured relative to an internalbiphenyl standard with a signal at 7.45 ppm) is between 200 and 230ligands per QD at all points in the titration, see Table 9 (the errorsarise from the discrepancy between integrating the whole peak vs.2×integrating half the peak). We therefore estimate the maximum numberof bound oleate per PbS QD (which equals the total number of oleate perQD minus the number of free oleate with zero HT added) to be 190-220.This number is used to approximate the number of bound MHO per QD.

TABLE 9 The Number of Bound and Free Oleate (OA) per QD and the TotalNumber of Binding Sites on the Surface of PbS QDs. Eq. of HT added toNo. of Free No. of Bound Total No. of Total No. of PbS QDs OA/QD^(a)OA/QD^(a) OA/QD^(b) binding sites^(c)  0 HT  10 200-220 210-230 200-220200 HT 140 80 220 210 400 HT 160 40 200 190 800 HT 180-200 20 200-220190-210 ^(a)Calculated by integrating the signals of OA vinyl protons in¹H NMR spectra as described in the text. ^(b)Estimated as the sum of thenumber of free and bound OA per QD. ^(c)Calculated by subtracting thenumber of free OA (with no HT added) from the total number of OA per QDestimated from b.

We acquired ¹H NMR spectra for all the six water-soluble PbS QD samples(13.2 μM) after ligand exchange and concentration calibration, as wedescribed in the previous section, and used a sum of five Lorentzianfunctions to fit and quantify the bound (one broad peak centered at˜2.07 ppm) and free (sharp triplet centered at ˜2.03 ppm) MHA species aswell as the singlet impurity located at ˜2.08 ppm. We only observe abroad feature of MHO for all QD samples, with no clear distinctionbetween bound and free populations, which suggests that either all theMHO ligands are bound to the surface of QD, or there's a fast exchangebetween bound and free populations of MHO. The poor resolution ofspectra also made it difficult to integrate these peaks accurately. Wetherefore chose to approximate the upper limit of the number of boundMHO per QD using the technique described above.

AQ is stable in aqueous solutions with the presence of water-soluble PbSQDs. We prepared four samples of 2.63 μM MHA-capped PbS QDs with 6000equivalents of AQ/QD added. All the samples were stored under the sameprotection of N₂ with different treatments of light. ¹H-NMR spectra showthat: (i) there's no spontaneous reaction between MHA-capped PbS QDs andAQ in the dark, (ii) when exposed to light, PbS QDs and AQ undergo aphotochemical redox reaction and create a long-living paramagnetic AQradical anion, which could be inferred from the broadened peaks in NMRspectra, (iii) the broad features of AQ radical anion resharpen when thesample we expose to light beforehand is put back in the dark overnight,which may indicate a charge-recombination process between AQ radicalanions and oxidized QDs, and (iv) we observed no new peaks in NMRspectra for any of these samples. We conclude that there's nospontaneous reaction between AQ and water-soluble PbS QDs without light,and we therefore chose to store all the samples in the dark for fourhours to equilibrate before taking PL measurements in order to eliminatephotochemical reactions that can be driven by room light.

PbS QD-AQ complexes could reach equilibrium of adsorption in four hours.We performed a time-dependent PL study on a separately prepared sample,using the same set-up described in last section (2.63 μM MHA-capped PbSQDs with 6000 equivalents of AQ/QD added). We took 17 measurementsconsecutively with a 30-min interval, and our result shows that theintegrated PL intensity saturates in four hours after we add AQ to theQD sample, which indicates that the adsorption of AQ on the surface ofQD has reached equilibrium.

Transient Absorption Spectra as A Proof for Electron Transfer andExtraction of Globally-shared Intrinsic eT Rate for Each eT-active PbSQD-AQ Pair. Ultrafast Transient Absorption Spectroscopy. We split the2.5-mJ output of a commercial amplified Ti-sapphire laser (Solstice, 1kHz, 100 fs, Spectra Physics), and guided 95% to an optical parametricamplifier (TOPAS-C, Light Conversion) used to produce the pumpwavelength (850 nm) for sample excitation, and 5% to a commercial TAspectrometer (HELIOS, Ultrafast Systems) for use as the probe for TAexperiments with pump-probe delay times up to 3200 ps. Within thespectrometer, a single filament broadband continuum of probe wavelengthsfrom 900-1400 nm was generated in a 1.2 cm thick sapphire plate and thenpassed through a long-wave pass filter to isolate near-infraredwavelengths above 850 nm. The probe light was then split into sample andreference beams. We combined the sample beam with the pump light at thesample, which was contained in a 2-mm quartz cuvette. The pump spot sizewas expanded to at least twice the size of the probe spot to compensatefor any imperfections in translation stage alignment. The transmittedprobe signal was collected into an optical fiber and dispersed onto anarray detector. Dividing the signal from the sample beam by the signalfrom the reference beam allowed us to divide out any fluctuations in theprobe beam intensity during the experiment. The output differentialabsorption spectrum (ΔA) was obtained through active backgroundsubtraction of the ground state spectrum by chopping the pump at 500 Hz.The pump light was depolarized to prevent unintentional photoselectionso that measurements reflect only population dynamics, and we adjustedthe power to 100 μW to ensure an expectation value (<N>) of ˜0.2 in thefirst excitonic state of the QDs and avoid any multi-exciton effects.The solution was stirred with a magnetic stir bar to minimize localheating.

Microsecond Transient Absorption Spectroscopy. We used a commercialspectrometer (EOS, Ultrafast Systems) to collect TA spectra forpump-probe delay times from 0.5 ns-25 μs. The excitation beam wasgenerated by the same method and followed the same beam path as theexcitation used for the picosecond TA experiment described above. Theproprietary EOS light source generates a super-continuum (400-1700 nm)probe pulse by focusing a diode laser into a photonic crystal fiber. Therepetition rate of the probe pulse is 2 kHz, which is twice therepetition rate of the pump pulse, and it is triggered in sync with thepump pulse. The pump-probe delay is electronically generated andmeasured by an electronic timer-counter-analyzer (Pendulum). After theprobe pulse passes through an 850-nm long pass filter, it is split by abeam-splitter into sample and reference beams. The sample beam was sentonto the sample, which was contained in a 2-mm quartz cuvette, andfocused to a spot size of ˜200 μm. The reference and sample beams arethen collected in optical fibers and dispersed onto array detectors. Inorder to achieve reasonable signal to noise, the incident power of pumplight was adjusted to be 2 mW. The solution was stirred with a magneticstir bar to minimize local heating. We performed transient absorptionexperiments, as described above, on a set of separately prepared 6.58 μMPbS QDs with two different surface charge densities and various amountsof AQ added, which we listed in Tables 10 and 11. The samples wereallowed to equilibrate in dark for four hours before we tookmeasurements. We applied a sum of exponentials convoluted with aninstrument response function (in this case, a Gaussian pulse) to fit thekinetic trace of the ground state bleach and determine the life time andrelaxation dynamics of photo-excited electrons and holes on both theultrafast and the microsecond timescale, eq 7:

$\begin{matrix}{y = {{IRF} \otimes {\sum\limits_{1}^{n}{A_{n}e^{\frac{- {({t - t_{0}})}}{\tau_{n}}}}}}} & (7)\end{matrix}$

Here we choose n to be the minimal integer that can make the fittingconverge. FIG. 7 shows the microsecond TA kinetic traces of 115MHA-capped and 31 MHA-capped PbS QDs extracted at 1014 nm and 1039 nm,respectively, while the ultrafast TA kinetics are described in FIG. 5,main text. The fitting parameters of these kinetic traces are listed inTable 10 and 11.

TABLE 10 Time Constants for Decay of the QD Exciton for QD-AQ Complexeson Ultrafast Time Scale. No. of bound Eq. of MHA AQ τ₁ = No. of ligandsbound/ PL/ τ_(eT) (ps) τ₂ (ps) τ₃ (μs) Sample per QD QD^(a) PL₀(A₃)^(b,c) (A₂)^(b,d) (A₁)^(b,e) 1 ~115 0 1 — 793 ± 244 2.06 ± 0.04(−0.06) (−0.87) 0.60 0.55 576 ± 84 — 2.06 ± 0.04 (−0.11) (−0.82) 0.860.42 574 ± 56 — 2.06 ± 0.04 (−0.16) (−0.77) 6 ~31 0 1 — 590 ± 105 1.61 ±0.02 (−0.08) (−0.88) 2.68 0.07 129 ± 4  590 ± 105 1.61 ± 0.02 (−0.70)(−0.18) (−0.09) 4.42 0.01 102 ± 2  590 ± 105 1.61 ± 0.02 (−0.84) (−0.11)(−0.02) ^(a)Calculated from the normalized PL intensity (PL/PL₀) of thesame samples. ^(b)Error bars are fitting errors. ^(c)Corresponds to thecharge separation process (electron transfer from QD conduction band tothe LUMO of AQ). The rate of charge recombination (recombination of theelectron on AQ with the hole in the QD valence band) could not bedeconvolved from the component corresponding to the charge trappingprocess, τ₂. ^(d)Corresponds to an electron or hole trapping process.For QD samples with 31 bound MHA ligands, this time constant was fixedto the value extracted from the fit of the QD-only sample. For QDsamples with 115 bound MHA per QD, however, τ₁ and τ₂ could not bedeconvolved in samples with added AQ. ^(e)Obtained from fits of themicrosecond-timescale TA kinetic traces, and fixed to these values whenwe fit the picosecond-timescale TA kinetic traces.

TABLE 11 Time Constants for Decay of the QD Exciton for QD-AQ Complexeson Microsecond Timescale. No. of Eq. of MHA Eq. of AQ τ₁ (μs) τ₂ (μs)Sample bound/QD added/QD (A₁)^(a) (A₂)^(b) 1 ~115 0 2.06 ± 0.04 0.19 ±0.07 (−0.86) (−0.07) 4000 1.05 ± 0.03 0.08 ± 0.01 (−0.73) (−0.21) 6 ~310 1.61 ± 0.02 0.06 ± 0.01 (−0.83) (−0.13) ^(a)corresponds to theradiative lifetime of PbS QDs ^(b)corresponds to the surface-mediatedcharge trapping processes.

Conversion of No. of bound MHA to Interfacial Charge Density. Assumingthat the charged MI-1A ligands are evenly distributed on the surface ofa QD and the ligand shell is, therefore, spherically symmetric, we applyeq 8 to convert the average number of bound MHA to interfacial chargedensity (charges/nm²):

$\begin{matrix}{= \frac{\begin{matrix}{{interfacial}\mspace{14mu} {charge}\mspace{14mu} {density}\mspace{14mu} \left( {{charges}\text{/}{nm}^{2}} \right)} \\{{{No}.\mspace{14mu} {of}}\mspace{14mu} {bound}\mspace{14mu} {MHA}\mspace{14mu} {per}\mspace{14mu} {QD}}\end{matrix}}{{total}\mspace{14mu} {surface}\mspace{14mu} {area}\mspace{14mu} {of}\mspace{14mu} a\mspace{14mu} {charged}\mspace{14mu} {ligand}\mspace{14mu} {shell}}} & (8)\end{matrix}$

The total surface area of a charged QD ligand shell can be calculatedusing eq 9, where

total surface area =4×π×[r(QD)+d(MHA)]²  (9)

r(QD)=1.6 nm is the radius of the inorganic core of a QD, and d(MHA)=1.3nm¹³ is the reported length of the MHA ligand. The calculatedinterfacial charge densities as well as the representative yields of eTupon mixing a 2.63 μM QD sample with 200 eq. of AQ are listed in Table12.

TABLE 12 Conversion of No. of bound MHA to Interfacial Charge Densityand the Correponding Yield of eT upon Adding 200 Eq. of AQ to a 2.63 μMQD sample. No. of Interfacial Yield of eT No. of bound MHA ChargeDensity upon Adding 200 Sample per QD^(a) (charges/nm²)^(b) Eq. ofAQ^(c) 1 115 ± 9  1.09 9.5 2 100 ± 8  0.95 6.1 3 94 ± 8 0.89 12.0 4 81 ±7 0.76 16.8 5 67 ± 6 0.63 50.7 6 31 ± 3 0.29 97.6 ^(a)Equal to theaverage number of bound MHA ligands per QD listed in Table 1.^(b)Calculated using eqs S12 and S13. ^(c)Extracted from FIG. 5B. Thelower yield of eT for sample 2 compared to sample 1 (presumably causedby experimental error) is against the general trend of our data, butthis discrepancy between sample 1 and 2 no longer shows up as weincrease the concentration of AQ.Reversible Modulation of the Electrostatic Potential of a ColloidalQuantum Dot through the Protonation Equilibrium of its Ligands

We demonstrate a precise method to control the photoluminescence (PL)intensity of a colloidal quantum dot (QD) through the acidity/basicityof its environment, using protons to regulate the permeability of theQD's ligand shell to a charged molecular quencher. The near-IRlight-emitting PbS QDs are passivated by an adlayer ofhistamine-derivatized dihydrolipoic acid (DHLA-His, FIG. 8). Theimidazole moieties at the outer boundary of this ligand shell capturefreely diffusing protons and switch from a neutral to positively chargedstate, such that the electrostatic field at the QD/solvent interface iscontrollable through the acidity of the solution. 1H NMR spectroscopyyields the average degree of protonation of the QDs' ligand shells, andsteady-state and time-resolved optical spectroscopies allow us to relatethis degree of protonation to the permeability of the ligand shell to acharged small-molecule electron acceptor, 9,10-anthraquinone-2-sulfonate(AQ), through the yield and rate of photoinduced electron transfer (e71)from the QD to AQ.

We previously described control over the electrostatic potential of acolloidal QD and its PL in the presence of a charged quencher withcharged, covalently bound ligands, and then more precisely with theseligands plus noncovalently bound counterions. Our introduction here of amechanism to reversibly tune the PL intensity of a QD via theprotonation equilibrium of its ligands suggests additionalfunctionality, specifically sensing of dynamic properties of a chemicalenvironment with a changing proton concentration, which could involvelocal proton fluxes or vesicle formation and expulsion. TheDHLA-functionalized PbS QD system is especially translatable to in vitrobiological applications because the emission of PbS QDs is tunablethroughout biological windows I and II (650-1350 nm; the QDs used hereemit at ˜1050 nm), and because the imidazolium group on DHLA has aphysiologically relevant pK_(a) (˜7.0) in water.

DHLA has a larger binding affinity for divalent metal cations and betterstability under acidic conditions (down to pH 3) than its monothiolateanalogues. We synthesized DHLA-His molecules via a published peptidecoupling reaction. We synthesized oleate-capped PbS QDs using an adaptedliterature procedure and prepared PbS QDs capped by a monolayer ofDHLA-His by adding ˜400 equiv of DHLA-His ligands per QD to a 2-mLsample of 400 μM oleatecapped PbS QDs dispersed in degassed hexanes andshaking the mixture gently until the QDs flocculated. We diluted thereaction mixture with 4 mL of degassed methanol and then centrifuged itat 7000 rpm for 5 min to precipitate the QDs. We washed the pellet with6 mL of ethyl acetate twice by centrifuging the suspension at 7000 rpmfor 5 min and decanting the clear supernatant. The cleaned QDs wereredispersed in 4 mL of degassed ethylene glycol (for opticalcharacterization) or methanol-d4 (for NMR measurements), and anyinsoluble QD aggregates were separated by centrifuging this solution at7000 rpm for 10 min. We collected the optically clear supernatant andused it as the stock solution for subsequent experiments. No boundoleate was detected on the surfaces of these QDs.

FIG. 9A shows the aromatic regions of ¹H NMR spectra of a series of0.132 mM solutions (in methanol-d4) of DHLA-His-capped PbS QDs mixedwith increasing molar equivalents (0-200 per QD) of p-toluenesulfonicacid (Tol-SO3H, pKa≈−3 inwater21). With no acid added (black trace atthe bottom), we observe (i) two sharp singlets located at ˜7.7 ppm (forHa) and ˜6.9 ppm (for Hb), which correspond to the aromatic protons onfreely diffusing DHLA-His molecules in solution, and (ii) two broadfeatures located at ˜7.6 ppm (Ha) and ˜6.8 ppm (Hb), which correspond tothe same protons on DHLA-His molecules that are bound to the surface ofQDs. Assignment of these two (bound and free) populations of DHLA-Hismolecules has been confirmed by NMR spin-spinmediated relaxation (T2)measurements. Each QD has an average of 76±8 DHLA-His ligands bound toits surface, and the ratio between freely diffusing and surfaceboundDHLA-His species is consistently 0.15±0.02. As we add increasing amountsof Tol-SO₃H to the QD dispersion, the protonation of the imidazole leadsto a more pronounced electron-withdrawing effect on the aromatic ringand results in a progressively downfield shift in the signals of thesetwo aromatic protons in both surface-bound and freely diffusing DHLA-Hisspecies FIG. 9A.

FIG. 9B is a plot of the peak position of H_(a) (in ppm), as a functionof Tol-SO₃H added, for both the bound (black; lower trace) and free(red; upper trace) populations of DHLA-His. The chemical shift of thesepeaks is a weighted average of the chemical shifts of the fullyprotonated (δ_(p)) and fully deprotonated (δ_(dp)) DHLA-His species.Assuming that (i) DHLA-His is 100% deprotonated before we introduce anyTol-SO₃H to the system, such that 6_(dp)=7.63 ppm, and (ii) DHLA-His is100% protonated as the chemical shift saturates (the equilibriumconstant for protonation of imidazole groups by Tol-SO₃H is ˜1010), suchthat δ_(p)=8.71 ppm (marked by the black dashed line in FIG. 9B), thedegree of protonation, p (0≤p≤1), for the imidazole moieties within theQD ligand shell is given by eq 10. We use eq 10 to translate thechemical shift of the aromatic proton signal

$\begin{matrix}{p = {{\frac{\delta_{obs} - \delta_{dp}}{\delta_{p} - \delta_{dp}} \times 100\%} = {\left( {{0.926 \times \delta_{obs}} - 7.06} \right) \times 100\%}}} & (10)\end{matrix}$

to the percentage of protonated imidazolium groups at the QD surface,FIG. 9B (inset).

In all experiments that follow, we limit the amount of Tol-SO₃H to 75equiv per QD because when we add more than 75 equiv DHLA-His ligandsbegin to desorb from the surface of the QD (presumably due to theprotonation of dithiolate anchoring groups).

We then split a batch of 6.58 μM QDs into four samples in ethyleneglycol and added either 0, 25, 50, or 75 mol equiv of Tol-SO₃H to eachsample to achieve a different concentration of protons at theligand/solvent interface (p=0, 31, 64, or 92%, respectively; see FIG.9B). We split each of these four samples into seven samples, to which weadded increasing molar equivalents (0-50 per QD) of AQ, a negativelycharged electron acceptor with respect to photoexcited PbS QDs. 1,2 Aswe have shown previously, 28-32 the probability of eT from thephotoexcited QD to a negatively charged AQ reflects the number of AQswithin a few Ångstroms of the QD surface, in the form of eitherstatically bound or transiently associated adsorbates. The photoinducedeT process results in a charge separated state that undergoesnonradiative recombination; therefore, the degree to which the PL of QDsis quenched in the presence of AQ is a sensitive measure of thepermeability of the QD ligand shell to the molecular probe, here,negatively charged AQ. We allowed all 28 samples to equilibrate in thedark (to avoid any potential photochemistry 1,33,34) under N2 overnightbefore performing any optical measurements. We then collected the PLspectra (850-1350 nm) of the samples in a 2 mm/10 mm dual path lengthquartz cuvette, photoexcited along the 10 mm axis at 800 nm.

FIG. 10A is a plot of “PL/PL0”, the integrated PL intensity for eachQD-AQ mixture (“PL”) divided by the integrated PL intensity of the samesample with no added AQ (“PL0”) vs the equiv of AQ added to the QDs forfour series of QDs with different degrees of protonation (p). For eachseries of QDs, the yield of PL quenching□that is, the yield ofeT□increases as we increase the molar equivalents of AQ added, asexpected (FIG. 10A, inset). The key result is that as p increases from 0to 92% the yield of eT is progressively enhanced for a given amount ofAQ added. This change is not due to the shift in the reduction potentialof AQ as a function of proton concentration in the solution (the drivingforce for eT from the QD to AQ changes negligibly) or to irreversiblephotooxidation of QDs, as evidenced by cyclic voltammetry and continuousillumination experiments. This enhancement is therefore causedexclusively by an increase in the average number of AQs adsorbed,statically or transiently, to each QD, due to enhanced permeability ofthe increasingly positively charged ligand shell to negatively chargedAQ.

To support this conclusion, we performed a series of transientabsorption (TA) measurements on QD solutions and QD-AQ mixtures([AQ]/[QD]=10:1) titrated with 0 (p=0), 25 (p=31%), or 75 (p=92%) molarequiv of Tol-SO₃H. The eT and subsequent charge recombination (CR),evidenced by an acceleration of the recovery of the ground-state bleachof the QDs upon addition of AQ, occur on both picosecond-to-singlenanosecond (FIG. 10B) and nanosecond-to-microsecond (FIG. 10C) timescales. We assign the dynamics on the faster time scale to eT and CRwithin statically bound QD-AQ complexes and the dynamics on the slowertime scale to the diffusion-controlled, collision-gated eT processwithin transiently associated QD and AQ species. The kinetic traces inFIGS. 10B and 10C show that addition of increasing amounts of Tol-SO₃Hin the presence of a fixed amount of AQ progressively increases the rateof eT on both time scales that we monitor, while addition of Tol-SO₃Halone (no AQ) has negligible impact on the exciton dynamics of the QD.These results are consistent with the trend of increasing PL quenchingupon progressive protonation of the QD ligand shell.

We determined the time constants for eT within static QD-AQ complexes byglobally fitting the picosecond-to-nanosecond kinetic traces in FIG. 10Bwith eq 11

ΔA(t)=IRF⊗[−A _(CS) e ^(−λ)(exp(λe ^(−t/τ) ^(Cs,static) )−1) −A_(CR)(1−e ^(−λ))e ^(−t/τ) ^(CR,static) −e ^(−λ) e ^(−t/τ) ¹ ]  (11)

where IRF is the instrument response function of the laser, τCS,staticis the time constant for charge separation between a photoexcited QD anda single absorbed AQ, τCR,static is the time constant for CR of the ionpair resulting from that charge separation, ACS and ACR account for therelative contributions of electrons and holes, respectively, to thetotal amplitude of the bleach signal, λ is the Poisson-averaged numberof statically adsorbed AQs per QD, and τ1 is the lifetime of excitonswithin QDs that have no statically adsorbed AQs. In fitting the kinetictraces in FIG. 10B, we fixed the values of τ1 to those measured by thelonger time scale TA experiment (see FIG. 10C and Table 13) because theycannot be measured accurately within the time window of ˜3 ns. We alsofixed the value of λ for each trace to λ=−ln(B/B0), where B/B0 is thefraction of QDs with zero adsorbed AQ molecules, measured from theamplitude of the ground-state bleach at delay times of 1800-3200 ps(well after the CR process within static QD-AQ complexes).

TABLE 13 Charge Separation and Recombination Dynamics for PbS QD-AQComplexes with Different Degrees of Protonation in Their Ligand ShellsτCS, static τCR, static avg. no. of Sample (ps) (ACS) (ps) (ACR)^(a)AQ/QD, λ^(b) τ1 (ns) (A1) τ2 (ns) (A2) τ3 (μs) (A3) avg. (μs)^(d) PbSQDs (p = 0) + 0.44  12 ± 0.6 240 ± 88  1.1 ± 0.07 0.87 10 equiv AQ(−0.09 ± 0.02)  (−0.15 ± 0.05) (−0.76 ± 0.05)  PbS QDs (p = 31%) + 9 ± 1240 ± 12  0.93 5.4 ± 0.2 110 ± 29 0.79 ± 0.09 0.30 10 equiv AQ (0.11 ±0.01) (0.29 ± 0.01) (−0.44 ± 0.03)  (−0.22 ± 0.03) (−0.34 ± 0.04)  PbSQDs (p = 92%) + 1.4 4.2 ± 0.2 100 ± 42 0.70 ± 0.34 0.12 10 equiv AQ(−0.65 ± 0.05)  (−0.22 ± 0.06) (−0.13 ± 0.06)  ^(a)These fittingparameters are shared among all three kinetic traces. Error bars arefitting errors from the kinetic traces extracted (at the ground-statebleach) from the picosecond-to-nanosecond TA spectrum of each sample,FIG. 10B. ^(b)The value of λ is fixed for each kinetic trace to thevalue of −ln(B/B0) for that trace, as explained in the text. ^(c)Errorbars are fitting errors from the kinetic traces extracted (at theground-state bleach) from the nanosecond-to-microsecond TA spectrum ofeach sample, FIG. 10C. ^(d)Amplitude-weighted averages of τ1, τ2, andτ3.

To first approximation, that is, if we assume that the protonation stateof the ligand shell only influences the number of AQ molecules thatadsorb to each QD surface and not the electronic coupling or drivingforce for eT between the QDs and these adsorbed acceptors□the value of pinfluences λ but not the intrinsic, single-donor, single-acceptor rateconstants τCS,static and τCR,static. We indeed find that λ, calculatedas described above, increases from 0.44 to 1.4 AQs adsorbed per QD as pincreases from 0 to 92%, Table 13. Fixing these values of λ and sharingthe values of τCS,static and τCR,static in fits of all three traces inFIG. 10B with eq 11, we find τCS,static=9 ps and τCR,static=240 ps.These rates are faster, by up to a factor of 5, than those we measuredfor the same acceptor but larger QDs as donors (radius of 1.6 nm insteadof 1.3 nm). The difference is reasonable since the driving force for eTin our current system is ˜0.3 eV larger than that for the previoussystem. The protonation state of the ligand shell should also increasethe average rate of diffusion-controlled eT from the QD to AQ byincreasing the diffusion constant of AQ within the ligand shell. Weobserve this increase by fitting the nanosecond-tomicrosecond kinetictraces that describe diffusion-controlled eT dynamics (FIG. 10C) with asum of three exponential functions convoluted with the IRF to obtain thetime constants listed in Table 13. The increasing rates of exciton decayon single-to-tens-of-ns (τ1), hundreds-of-ns (τ2), and single-μs (τ3)time scales upon progressive protonation of the QD ligand shell suggestthat attractive forces between the ligand shell and AQ enhance the rateof eT through a continuum of diffusion-controlled pathways.

Finally, we demonstrate acidity/basicity-controlled reversible cyclingof the permeability of the QD's ligand shell. We prepared two 3 mL, 10.0μM samples of DHLA-His-capped PbS QDs and pretreated one of the sampleswith 10 equiv of AQ, while the other sample served as the “blank”control. We then alternately mixed 75 equiv of Tol-SO₃H and 75 equiv oftetramethylammonium hydroxide (NMe₄OH) into each sample to cycle theligand shell between protonated and deprotonated states and collectedthe steady-state PL spectra immediately after each addition, FIG. 11.The protonation/deprotonation process takes place instantaneously. Whenno AQ is added, the PL intensity of the QD ensemble remains stable uponsequential additions of acid/base (FIG. 11, red trace), which suggeststhat 75 equiv of acid/base alone does not impact the surface passivationof these QDs, consistent with the results from NMR. The PL intensity ofthe QDs however shows dramatic, reversible oscillation, with an averageratio of PL(p=0)/PL(p=92%) of 6.7 (a maximum observed ratio of 28),which confirms that both AQ and surface-bound DHLA-His ligands do notdegrade during the cycling of acid-base titrations, and demonstratesthat the QD-AQ system is a sensitive, robust optical probe of the protonconcentration in solution. We suspect that the slight enhancement in thePL intensity of these QDs, in both acidic and basic conditions, over thecourse of the cycling is due to the increasing ionic strength of thesolution, which lowers the probability that AQ will diffuse to the QDsurface by stabilizing it in bulk solution.

In summary, we have utilized the protonation equilibrium of the ligandson a PbS QD to reversibly modulate its PL and exciton lifetime. Byincreasing the positive charge density at the outer boundary of a PbS QDfrom 0 to 70 charges per QD, we decreased the PL of the QD sample by afactor of 6.7 and decreased the average exciton lifetime of the QDs by afactor of 7 by increasing the permeability of the QD's ligand shell to acharged molecular electron acceptor, AQ. These changes in electrostaticpotential are reversible through sequential additions of acid and baseto the QD dispersion. Our analysis treats the enhanced permeability ofthe QD ligand shell upon protonation exclusively as an electrostaticeffect and neglects possible contributions from changes in themorphology of the ligands upon their protonation. We believe that ourfocus on electrostatics is justified, however, because most analyses ofthese morphological changes predict that the permeability of themonolayer should increase (due to formation of patches, ridges, or thinregions) as the degree of protonation decreases due to competingelectrostatic and van der Waals interactions. This trend is the oppositeof what we see.

This work also demonstrates the use of an interfacial redox process as ahighly sensitive probe for the local chemical and electrostaticenvironment of an emissive nanoparticle. Our results suggestapplications in not only steady state but also dynamic proton sensing,with a response time limited by diffusion of the quencher to the surfaceof the QD, a process that could be sped up by tethering it to the QDsurface with a flexible linker. We could also measure the pH of anaqueous system by first experimentally determining the pK_(a) of theimidazolium group in the bound DHLA-His molecules (via NMR,potentiometric, or spectrophotometric titration of the sample with anacid or base of known pK_(a)) and then translating the degree ofprotonation within the ligand shell, p, into the concentration of freelydiffusing protons in solution. Furthermore, we could accomplish sensingof other ions by incorporating specific binding sites into the QD'sligands.

Synthesis of Histamine-Derivatized Lipoic Acid (LA-His) andHistamine-Derivatized Dihydrolipoic Acid (DHLA-His). We synthesizedhistamine-derivatized lipoic acid (LA-His) using a well-establishedpeptide coupling reaction, see Scheme 1. We dissolved 4.5 g (21.8 mmol,1 equiv.) of lipoic acid, 4.0 g (21.8 mmol, 1 equiv.) of histaminedihydrochloride, and 15 mL (87.0 mmol, 4 equiv.) ofN,N-diisopropylethylamine (DIPEA) in 75 mL N,N-dimethylacetamide. Wethen initiated the coupling reaction by adding 11.5 g (26.1 mmol, 1.2equiv.) of benzotriazole-1-yl-oxy-tris-(dimethylamino)-phosphoniumhexafluorophosphate (“BOP reagent”) to the solution, and allowed thereaction to proceed under room temperature for at least 60 hours. Thereaction mixture was then diluted with 200 mL deionized water and theproduct was extracted six times with ethyl acetate (100 mL perextraction). The organic layer was dried over anhydrous sodium sulfate,and the filtrate was concentrated under reduced pressure andchromatographed on a silica gel column using CH₂Cl₂/CH₃OH (9:1) aseluent. Yield: 2.2 g (34%); R_(f)=0.5 (CH₂Cl₂/CH₃OH (9:1)). ¹H NMR(Methanol-d₄, 600 MHz): δ 7.80 (s, 1H), 6.93 (s, 1H), 3.55 (dq, J=9.0Hz, 6.3 Hz, 1H), 3.43 (t, J=7.1 Hz, 2H), 3.19-3.06 (m, 2H), 2.79 (t,J=7.1 Hz, 2H), 2.45 (dq, J=12.5 Hz, 6.4 Hz, 1H), 2.17 (t, J=7.4 Hz, 2H),1.87 (dq, J=13.5 Hz, 6.8 Hz, 1H), 1.74-1.43 (m, 4H), 1.49-1.33 (m, 2H);MS-ESI: m/z=300.16 ([M+H]⁺, predicted: 300.46), 322.15 ([M+Na]⁺,predicted: 322.45).

We prepared DHLA-His by reducing LA-His with NaBH₄, a procedurewell-described in the literature [Uyeda et al., Synthesis of CompactMultidentate Ligands to Prepare Stable Hydrophilic Quantum DotFluorophores. J. Am. Chem. Soc. 2005, 127, 3870-3878; Liu et al.,Compact Biocompatible Quantum Dots Functionalized for Cellular Imaging.J. Am. Chem. Soc. 2008, 130, 1274-1284; Muro et al., Small and StableSulfobetaine Zwitterionic Quantum Dots for Functional Live-Cell Imaging.J. Am. Chem. Soc. 2010, 132, 4556-7]. In a typical synthesis, we firstdissolved 2.2 g (7.5 mmol, 1 equiv.) LA-His with 40 mL methanol in a500-mL round-bottom flask, and cooled down the solution with an icebath. We added 1.13 g NaBH₄ (30.0 mmol, 4 equiv.) slowly to thissolution under constant stirring. After 30 min, we diluted the colorlessreaction mixture with 200 mL deionized water, and extracted the productsix times with ethyl acetate (100 mL per extraction). The organic layerwas dried over anhydrous sodium sulfate, and the solvent was evaporatedunder reduced pressure to yield the product as a clear, colorless oil.Yield: 2.0 g (90%). ¹H NMR (Methanol-d₄, 600 MHz): δ 7.58 (s, 1H), 6.84(s, 1H), 3.41 (t, J=7.2 Hz, 2H), 2.91-2.84 (m, 1H), 2.77 (t, J=7.2 Hz,2H), 2.72-2.60 (m, 2H), 2.17 (t, J=7.4 Hz, 2H), 1.92-1.84 (m, 1H),1.75-1.35 (m, 7H); MS-ESI: m/z=302.06 ([M+H]⁺, predicted: 302.47).

Synthesis and Purification of Oleate-capped PbS QDs. We synthesizedoleate-capped PbS QDs with a radius of ˜1.3 nm and a first excitonicpeak at 800-860 nm using an adapted literature procedure [Hines andScholes, Colloidal PbS Nanocrystals with Size-Tunable near-InfraredEmission: Observation of Post-Synthesis Self-Narrowing of the ParticleSize Distribution. Adv. Mater. 2003, 15, 1844-1849]. Briefly, we mixed0.36 g PbO and 1.0 mL oleic acid with 19.0 mL 1-octadecene (ODE) in a50-mL three-neck round bottom flask at room temperature. Heating themixture up to 150° C. with constant stirring under N₂ flow for an hourproduced a dear and colorless solution. We cooled the mixture to 110°C., and injected 0.17 mL of hexamethyldisilathiane dissolved in 8 mL ofdegassed ODE. The solution turned from orange to brown within 3 seconds.After 4 minutes, we remove the heating mantle and allow the reactionmixture to cool down naturally to room temperature. The product wasseparated into four 50-mL centrifuge tubes for further purification. Wepurified the QDs by first washing the reaction mixture with acetone (6:1by volume), and centrifuging it at 3500 rpm for 5 min. We then decantedthe supernatant, dried the QD pellet, redispersed the QDs in 5 mLhexanes, and precipitated the QDs two additional times, as describedabove, using 12.5 mL methanol and acetone, respectively, as thenon-solvents. The purified PbS QDs were finally dispersed in 20 mL ofhexanes to form the stock solution.

Assignment of Surface-bound and Freely Diffusing DHLA-His Species viaSpin-spin-mediated Relaxation (T₂) Measurements. We recorded ¹H NMRspectra of a 0.132 mM sample of DHLA-His-capped PbS QDs, pretreated with75 equiv. of p-tolunenesulfonic acid (Tol-SO₃H), on an Agilent DD2600-MHz NMR spectrometer. We chose a spin-echo pulse sequence for T₂relaxation measurement, set the relaxation delay between scans to be 10s, and arrayed the dephasing time, t, between 0.01 and 6.4 s. Figure S2shows two kinetic traces of T₂ decay of the H_(a) proton signal (seeFIG. 9A). Each kinetic trace fits to a single-exponential function, eq12.

I=I ₀ ×e ^(−t/T) ²   (12)

The T₂ lifetime of the broad feature (black trace) is shorter than thesharp feature (red trace) by a factor of 7 (1.2 s vs. 0.18 s). This moreefficient transversal relaxation for the broad Ha signal ispredominantly caused by the restricted rotational mobility of DHLA-Hisligands that are bound to the surface of QDs, and has been observed byboth our group⁸ and others⁹. We therefore assign the sharp and broadfeatures in FIG. 9A to freely diffusing and surface-bound ligands,respectively.

Quantification of DHLA-His Ligands within the Ligand Shell of PbS QDs.We prepared DHLA-His-capped PbS QDs using the procedures described inthe main text, and applied ¹H NMR to quantify the number of boundDHLA-His ligands per QD. The NMR samples were 0.132 mM solutions of QDsin methanol-d₄, with 100 equiv. of sodium formate as an internalintegration standard (singlet at 8.56 ppm, 1H). Experiments wereperformed on an Agilent DD2 600-MHz NMR spectrometer. We set theacquisition time to 27 s and the relaxation time to 90 s, respectively,to allow for complete collection of the free induction decay signal andsufficient relaxation of proton nuclei between measurements, andperformed 8 scans to get a spectrum with satisfactory signal-to-noiseratio. The 6.5-7.1 ppm region of the resulting spectra, which containsthe aromatic proton signal (H_(b), see FIG. 9A) of DHLA-His, is fit witha sum of two Lorentzian functions, and the broad featurecentered at ˜6.8ppm, which corresponds to those protons of surface-bound DHLA-Hisligands, is integrated against the sodium formate internal standard. Thefitting results of ¹H NMR spectra were acquired from three separatelyprepared samples. There are, on average, 76±8 DHLA-His ligands bound tothe surface of each QD.

Full NMR Spectrum of DMA-His-capped PbS QDs. The full NMR spectrumcollected from a methanol-d₄ solution of 0.132 mM DHLA-His-capped PbSQDs (data not shown). Sharp peaks and broad features were buriedunderneath correspond to DHLA-His ligands that are freely diffusing orbound to the surface of QDs, respectively. The absence of features at˜5.5 ppm indicates that all the native oleate ligands are displacedduring the ligand exchange.

DHLA-His ligands begin to desorb from the surface of QD upon mixing with≥100 equiv. of Tol-SO₃H. We calculate the ratio between freely diffusingand surface-bound DHLA-His ligands through dividing the area of thesharp singlet (H_(a), see FIG. 9A) by the area of the correspondingbroad feature. FIG. 13 contains a plot of this ratio as a function ofthe equiv. of acid we mix into the QDs. When 0-75 equiv. acid was added,this ratio remains relatively constant (0.16±0.04); when more than 100equiv. of acid was added instead, however, this ratio starts to increasedramatically, as we described in the main text, presumably due to theprotonation and desorption of the surface-bound DHLA-His molecules. Wetherefore limited the equiv. of acid up to 75 equiv. for all ourphotophysical characterizations to make sure that QDs with differentdegrees of protonation at their ligand shells have a comparable coverageof ligands on their surface.

The PL intensity of the QDs saturates within 2 h upon mixing with AQ. Weprepared a 3-mL, 10 μM ethylene glycol solution of DHLA-His-capped PbSQDs, contained in a 1 cm×1 cm air-tight quartz cuvette and pretreatedwith 10 equiv. of AQ, and performed a time-dependent PL study using thesame set-up described in the main text. We took 10 measurementsconsecutively, and our result (see FIG. 14) shows that the integrated PLintensity saturates within two hours of adding AQ to the QD sample,which indicates that the adsorption of AQ on the surface of QD hasreached equilibrium.

Ground-state Absorption Spectra of DHLA-His-capped PbS QDs withIncreasing Degree of Protonation in Their Ligand Shells. We superimposedground-state absorption spectra of 13.2 μM ethylene glycol solutions ofDHLA-His-capped. PbS QDs, pretreated with 0, 25, 50 or 75 equiv.Tol-SO₃H (FIG. 15). The peak position of these spectra stay constant,which suggests that these QDs do not degrade upon addition of up to 75equiv. acid, in line with our observation from NMR, see FIG. 13. Theslight decrease (<10%) in the absorbance, however, is likely caused bythe surface reconstruction upon addition of acid into the solution.

Continuous Illumination Experiments of DHLA-His-capped QDs. In order toprove that the enhanced PL quenching in FIG. 10A upon addition of acidis not a result of irreversible photooxidation of QDs by AQ in thepresence of protons, we prepared a 3-mL, 10.0 μM sample ofDHLA-His-capped PbS QDs, pretreated with 75 equiv. Tol-SO₃H and 5 equiv.AQ, in a 1 cm×1 cm air-tight quartz cuvette, and illuminated this samplewith a 5-mW, 532-nm laser pointer under constant stirring overnight.FIGS. 16A and 16B contains the ground-state absorption and PL spectra ofthis sample before and after the illumination. The negligible change inboth spectra over time suggests that QDs remain stable throughout theillumination experiment, and irreversible photooxidation is, therefore,not the cause of increased PL quenching.

Cyclic Voltammetry (CV). In order to determine whether, in our case, thereduction potential of AQ changes as the acidity of the solutionincreases, we performed cyclic voltammetry experiments using athree-electrode cell with a 3-mm radius glassy carbon electrode as theworking electrode, silver wire as the pseudo reference electrode, andplatinum wire as the counter electrode. Measurements were taken withPrinceton applied research potentiostats using 1 mM degassed ethyleneglycol solutions of AQ, with 0.25 M NH₄PF₆ added as the supportingelectrolyte and 2 mM ferrocene added as the internal standard.

The cyclic voltammogram in FIG. 17 shows the reduction of AQ under ascanning rate of 0.1 V/s. The observed reduction potential of AQ isconsistently ˜−0.8 V vs. Fc/Fc⁺ redox couple, as the concentration ofTol-SO₃H increases from 0 to 0.9 mM (the maximum concentration of acidwe used in our PL studies). We suspect that Tol-SO₃H cannot fullydissociate into freely diffusing protons and Tol-SO₃ ⁻ anions inethylene glycol, so its impact on the acidity of the solution may not beas dramatic as the case in water, which explains the insensitivity ofthe reduction potential of AQ to the concentration of acid added.Furthermore, we have approximately 70 imidazole groups per QD, and weadd a maximum of 75 equivalents of acid per QD, so in the presence ofQDs, protons are mostly adsorbed at the outermost boundary of the QDligand shell, in the form of imidazolium, instead of freely diffusing insolution as they would be in water, and the actual activity of protonsis, therefore, much smaller than in water.

We therefore conclude that the increasing yield of PL quenching that weobserve in FIG. 10A is not a result of the change in driving force foreT due to the presence of protons in solution, and can be treated as anelectrostatic effect.

Transient Absorption Measurements. Picosecond-to-nanosecond TransientAbsorption Spectroscopy. We split the 2.5-mJ output of a commercialamplified Ti-sapphire laser (Solstice, 1 kHz, 100 fs, Spectra Physics),and guided 95% to an optical parametric amplifier (TOPAS-C, LightConversion) used to produce the pump wavelength (810 nm) for sampleexcitation, and 5% to a commercial TA spectrometer (HELIOS, UltrafastSystems) for use as the probe for TA experiments with pump-probe delaytimes up to 3200 ps. Within the spectrometer, a single filamentbroadband continuum of probe wavelengths from 900 to 1400 nm wasgenerated in a 1.2-cm thick sapphire plate and then passed through along-wave pass filter to isolate near-infrared wavelengths above 850 nm.The probe light was then split into sample and reference beams. Wecombined the sample beam with the pump light at the sample, which wascontained in a 2-mm quartz cuvette. The transmitted probe signal wascollected into an optical fiber and dispersed onto an array detector.Dividing the signal from the sample beam by the signal from thereference beam allowed us to divide out any fluctuations in the probebeam intensity during the experiment. The output differential absorptionspectrum (ΔA) was obtained through active background subtraction of theground state spectrum by chopping the pump at 500 Hz. The pump light wasdepolarized to prevent unintentional photoselection so that measurementsreflect only population dynamics, and we adjusted the power to 100 μW toavoid any multi-exciton effects. The solution was stirred with amagnetic stir bar to minimize local heating.

Nanosecond-to-microsecond Transient Absorption Spectroscopy. We used acommercial spectrometer (EOS, Ultrafast Systems) to collect TA spectrafor pump-probe delay times from 0.5 ns-15 μs. The excitation beam wasgenerated by the same method and followed the same beam path as theexcitation used for the picosecond-to-nanosecond TA experiment describedabove. The proprietary EOS light source generates a super-continuum(400-1700 nm) probe pulse by focusing a diode laser into a photoniccrystal fiber. The repetition rate of the probe pulse is 2 kHz, which istwice the repetition rate of the pump pulse, and it is triggered in syncwith the pump pulse. The pump-probe delay is electronically generatedand measured by an electronic timer-counter-analyzer (Pendulum). Afterthe probe pulse passes through an 850-nm long pass filter, it is splitby a beam-splitter into sample and reference beams. The sample beam wassent onto the sample, which was contained in a 2-mm quartz cuvette, andfocused to a spot size of ˜200 μm. The reference and sample beams arethen collected in optical fibers and dispersed onto array detectors. Inorder to achieve reasonable signal to noise, the incident power of pumplight was adjusted to 2 mW. The solution was stirred with a magneticstir bar to minimize local heating.

We performed TA experiments, as described above, on a set of separatelyprepared ethylene glycol solutions of 13.2 μM PbS QDs. The compositionsof these samples are described in the main text. All samples wereallowed to equilibrate in the dark overnight before we took TAmeasurements. FIG. 18A contains a representative spectrum (extracted at2 ps) of our TA measurements. The kinetic traces are extracted at theground state bleach for all TA spectra. While the kinetic traces forps-to-ns TA were fit using eq 10 in the main text, the kinetic tracesfor ns-to-μs TA were simply fit with a sum of three exponentialsconvoluted with an instrument response function (IRF), see eq 13, sincethe rate constant for diffusion-controlled eT does not depend

$\begin{matrix}{{\Delta \; {A(t)}} = {{IRF} \otimes {\sum\limits_{n = 1}^{3}\; {A_{n}e^{\frac{- {({t - t_{0}})}}{\tau_{n}}}}}}} & (13)\end{matrix}$

directly on the number of AQs that simultaneously interact with each QD.

We also performed the same experiments on the same samples with no AQadded, which we used as the “blank” controls. FIGS. 18B and 18C containsthe kinetic traces (extracted at the ground-state bleach) of the controlsamples for both ps-to-ns and ns-to-μs TA. Our results show that theaddition of Tol-SO₃H has negligible impact on the exciton dynamics ofQDs: only up to 8% recovery, presumably due to surface-mediated chargetrapping processes, shows up in the kinetics of ps-to-ns TA ofQD—Tol-SO₃H mixtures, and completes within 5 ps, while the kinetictraces extracted from the ns-to-μs TA of QDs with different degrees ofprotonation satisfactorily overlay with each other. Here we simply fitthe ns-to-μs kinetic traces with a sum of two exponentials convolutedwith the IRF, eq 14 (see the respective fitting parameters in Table 14).

$\begin{matrix}{{\Delta \; {A(t)}} = {{IRF} \otimes \left\lbrack {{A_{1}e^{\frac{- {({t - t_{0}})}}{\tau_{1}}}} + {A_{2}e^{\frac{- {({t - t_{0}})}}{\tau_{2}}}}} \right\rbrack}} & (14)\end{matrix}$

TABLE 14 Fitting Parameters for the Kinetic Traces Extracted fromNano-to-Microsecond TA Spectra of PbS QDs with Different Degrees ofProtonation in Their Ligand Shells. Nanosecond-to-Microsecond TAKinetics τ₂ (μs) T₃ (μs) Sample (A₁)^(a) (A₂)^(a) PbS QDs + 0.16 ± 0.04 1.4 ± 0.04 0 equiv. acid 0.16 ± 0.04 (−0.89 ± 0.02) PbS QDs + 0.17 ±0.05  1.5 ± 0.03 25 equiv. acid (−0.11 ± 0.02)  (−0.89 ± 0.02) PbS QDs +0.19 ± 0.06  1.5 ± 0.06 75 equiv. acid (−0.15 ± 0.03)  (−0.85 ± 0.03)^(a)Error bars are fitting errors from the kinetic traces extracted (at901 nm) from the nanosecond-to-microsecond TA spectrum of each sample,FIG. 18C.

Calculations of the Amplitude-weighted Average of Excited-stateLifetimes. We applied eq 15 to calculate the amplitude-weighted averageof excited-state lifetime of the QD exciton using

$\begin{matrix}{\overset{\_}{\tau} = \frac{\Sigma_{i = 1}^{3}A_{i}\tau_{i}}{\Sigma_{i = 1}^{3}A_{i}}} & (15)\end{matrix}$

the time constants we measured by nanosecond to microsecond TA (seeTable 13).

Calculation of the Average Number of Statically Adsorbed AQs per QD, λ.As we and others have demonstrated in the previous work,₁₁₋₁₃ within ahomogeneous QD-AQ mixture, the probability of finding a QD within theensemble with n adsorbed AQ molecules, p (n), is well-described by aPoisson distribution, eq 16. Given that the charge separation (CS) andcharge

$\begin{matrix}{{p(n)} = {\frac{\lambda^{n}}{n!} \times e^{- \lambda}}} & (16)\end{matrix}$

recombination (CR) processes within statically bound QD-AQ complexescomplete in hundreds of ps (see the main text, Table 13), and assumingthat each static QD-AQ pair will necessarily participate in eT uponphotoexcitation (the rate of static eT typically outcompetes the rate ofradiative recombination by a factor of at least 4000₁₂), the fraction ofphotoexcited QDs that have no adsorbed AQ, i.e., the fraction of theseQDs that do not participate in static eT, should be equal to thefraction of the ground-state beach signal left at the end of thepicosecond-to-nanosecond TA, eq 17. We normalize the kinetic traceextracted at the ground-state bleach (878 nm) to its

p(n=0)=e ^(−λ) =B/B ₀  (17)

minimum, B₀, and average the bleach signal between the time delay of1800-3200 ps (well after CR) to obtain the value of B/B₀. The averagenumber of statically adsorbed AQs per QD, λ, could therefore becalculated using eq 18. We chose to fix the value of λ in our globalfits of the

λ=−ln(B/B ₀)  (18)

picosecond-to-nanosecond TA kinetics (see FIG. 10B and Table 13 in themain text) in order to constrain the fits and minimize the codependencebetween the parameters.

Time-dependent Study of the Acid/Base Cycling Experiment. FIG. 19contains a plot of the integrated PL intensity of the QD sample (3-mL,10.0 μM in ethylene glycol with 10 equiv. AQ added, see FIG. 11) as afunction of time after addition of acid/base. The PL intensity remainsrelatively constant over an hour for both cases, which suggests that theprotonation/deprotonation of the QD ligand shell and the resultingchange in its permeability to AQ takes place instantaneously.

The presence of electrolyte contributes to the slight enhancement in thePL of QDs. To test our hypothesis that the enhanced PL of QDs over thecycling experiment in FIG. 11 is caused by the increasing ionic strengthof the solution, we tracked the integrated PL intensity of two 3-mL,10.0 μM PbS QD samples (p=0 and p=92%, respectively, with 10 equiv. AQadded), mixed with increasing molar equivalents (0-300 per QD) oftetramethylammonium chloride (NMe₄Cl) salt, which we added 75 equiv. ata time, see FIG. 20. All the PL spectra are normalized to the PL of thesame sample with no AQ added. The plot clearly shows that, for bothfully protonated and fully deprotonated QDs, the PL intensity graduallyrecovers upon introducing an electrolyte at the relevant concentration.

Electrostatic-Based Proton and Acetylation Sensing Using Quantum Dots(QDs)

Many chemical/biological processes can be monitored by tracing thechange of the state of electrostatic charges. We developed a generalmethodology for designing quantum dot (QD) probes based on mediating thecharge transfer process via the electrostatic interaction between the QDsurface and the acceptors. The surface of this nanoprobe is covered withthree types of ligands: 1) passivating ligands that protect the QDsurface and make the sensor soluble in aqueous environment, 2) ligandsthat can vary in their electrostatic charge in response to changes ofchemical environment, 3) photoluminescence mediators that are anchoredto QDs via flexible linkers and are attracted/repelled by QDs dependingon their surface electrostatic potential. Based on this strategy, aprobe of either pH or acetylation (it works for both) was fabricatedusing mercaptoalkylamine-capped QDs by decorating their surface withfive equivalents of tethered electron acceptors, methyl propyl viologen(MPV²⁺) (FIG. 21 and FIG. 22). Specifically, the sensor is prepared from155 eq. polyethylene glycol thiol (PEG-thiol MW 2000), 80 eq.6-amino-1-hexanethiol or 1-amino-8-octanethiol, and 5 eq MPV-PEG-thiol(MW 2000).

The change of the pH in the solution induces theprotonation/deprotonation of the amine groups and varies theelectrostatic potential of the surface. The electrostatic interactionbetween the MPV²⁺ and the QD surface determines the average distancebetween them, and hence governs the rate of photoinduced electrontransfer from the QD to MPV²⁺, which mediates the intensity of thephotoluminescence of QDs. The same sensor can also respond to theacetylation reaction of the amine groups on the QD surface. Titratingthe QD sensors with sulfo-NHS-acetate in a MOPS buffer solution (pH=7.2)diminishes the PL intensity because the acetylation reaction blocks thepositively-charged amine groups (FIG. 21 and FIG. 23). By replacing themercaptoalkyl amine ligands with peptides substrates which mimic theproteins in a biological system that undergo acetylation, these probescan be used to monitor the real-time level of protein acylation in vivo.

Electrostatic-Based Proton Sensing Using Quantum Dots (QDs)

We coat QDs using a mixed monolayer of dithiolate ligands withpH-responsive imidazole and electron-accepting rhodamine B (RhB)moieties, see FIG. 24. Upon protonating/deprotonating the pH-responsiveimidazole groups (pK_(a)=7 in water), we modulate the electrostaticpotential at the surface of QDs and, therefore, thedesorption/adsorption of the electron-accepting RhB moieties, whichdetermines the fluorescence intensity of the QD ensemble as a functionof the pH of the solution. The response time of this process canpotentially bypass the limit of diffusion (which is on the order of tensof nanosecond to microsecond), since the electron acceptors arecovalently tethered to the surface of QDs. Modulation of thefluorescence intensity of a PbS QD via tuning the electrostaticpotential at the ligand shell-solvent interface. The third component ofthe QD ligand shell, which consists of mercaptopolyethylene glycol 2000or polyethylene glycol-2000-derivatized dihydrolipoic acid, solubilizesthe QDs in water, and is omitted for clarity.

We claim:
 1. An environmental sensor comprising: (a) a photoluminescencequencher and (b) a nanoparticle comprising a photoluminescent core andplurality of reactive ligands bound to the core and forming a quencherpermeable ligand shell surrounding the core wherein each of the reactiveligands comprises a reactive moiety capable of being modulated between afirst state and a second state and an anchoring group for binding theligand to the core, and wherein the permeability of the ligand shell isdetermined by the proportion of the reactive ligands in a first stateand a second state.
 2. The sensor of claim 1, wherein the nanoparticlefurther comprises a diluent ligand, a solubilizing ligand, or both thediluent ligand and the solubilizing ligand.
 3. The sensor of claim 1,wherein the reactive moiety comprises an anionic charge in the firststate and a neutral charge in the second state.
 4. The sensor of claim1, wherein the reactive moiety comprises a cationic charge in the firststate and a neutral charge in the second state.
 5. The sensor of claim1, wherein the reactive ligand comprises a radical of formula A-T-R,wherein A comprises the anchoring group and the anchoring group isselected from the group consisting of a alkylmonothiolate, aalkyldithiolate, or an alkyl trithiolate, wherein R comprises thereactive moiety selected from the group consisting of a carboxyl, ahydroxyl, a sulfo, a sulfhydryl, a phosphoryl, a phosphate and aconjugate base thereof or a reactive moiety selected from the groupconsisting of a substituted or unsubstituted amine or alkylamine, asubstituted or unsubstituted imidazole, a substituted or unsubstitutedbenzimidazole, a substituted or unsubstituted pyrimidine, a substitutedor unsubstituted purine, a substituted or unsubstituted pyridine, asubstituted or unsubstituted pyrrolidine, and a conjugate acid thereof,and wherein T comprises a tether comprising —(CH₂)_(n)— where n is aninteger and n=1-15, —(CH₂)_(n)(CONH)(CH₂)_(m)— where n and m areintegers and n+m=1-15, or —(OCH₂CH)_(n)— where n is an integer andn=1-100.
 6. The sensor of claim 1, wherein the sensor further comprisesa flexible tether for tethering the quencher to the nanoparticle.
 7. Thesensor of claim 1, wherein the quencher comprises an anionic charge orcationic charge.
 8. The sensor of claim 1, wherein the quenchercomprises 9,10-anthraquinone-2-sulfonate, rhodamine B, or methyl propylviologen.
 9. The sensor of claim 1, wherein the quencher comprises aradical of formula A-T-Q, wherein A comprises the anchoring group andthe anchoring group is selected from the group consisting of aalkylmonothiolate, a alkyldithiolate, or an alkyl trithiolate, wherein Qcomprises an aromatic moiety capable of accepting an electron from thecore, and wherein T comprises a tether comprising —(CH₂)_(n)— where n isan integer and n=1-15 —(CH₂)_(n)—, —(CH₂)_(n)(CONH)(CH₂)_(m)— where nand m are integers and n+m=1-15, or —(OCH₂CH₂)_(n)— where n is aninteger and n=1-100.
 10. The sensor of claim 9, wherein the aromaticmoiety comprises 9,10-anthraquinone-2-sulfonate, rhodamine B, or methylpropyl viologen.
 11. The sensor of claim 1, wherein the first state is afirst protonation state and the second state is a second protonationstate.
 12. The sensor of claim 1, wherein the first state is anacetylated state and the second state is a deacetylated state.
 13. Anenvironmental sensor comprising: (a) a photoluminescence quencher and(b) a nanoparticle comprising a photoluminescent core and plurality ofcharged ligands bound to the core and forming a quencher permeableligand shell surrounding the core wherein each of the charged ligandscomprises a charged moiety capable of being modulated between a firststate and a second state and an anchoring group for binding the ligandto the core, and wherein the permeability of the ligand shell isdetermined by the proportion of the ligands in a first state and asecond state.
 14. The sensor of claim 13, wherein the first state is afirst protonation state and the second state is a second protonationstate.
 15. The sensor of claim 13, wherein the first state is anacetylated state and the second state is a deacetylated state.
 16. Thesensor of claim 13, wherein the first state is an ion-paired state andthe second state ion-unpaired state.
 17. A method for environmentalmonitoring, the method comprising: (a) contacting an environmentalsensor with a molecular environment, the sensor comprising: (i) aphotoluminescence quencher and (ii) a nanoparticle comprising aphotoluminescent core and plurality of ligands bound to the core andforming a quencher permeable ligand shell surrounding the core whereineach of the ligands comprises a charged moiety or a reactive moietycapable of being modulated between a first state and a second state andan anchoring group for binding the ligand to the core, and wherein thepermeability of the ligand shell is determined by the proportion of theligands in a first state and a second state, (b) irradiating the sensor,and (c) detecting a photoluminescent signal.
 18. The method of claim 17,wherein the molecular environment comprises a reactant capable ofgenerating a product and wherein the product is capable of reacting withthe reactive moiety to modulate the reactive moiety between the firststate and the second state.
 19. The method of claim 18, wherein theproduct is H⁺, OH⁻, an acid, or a base.
 20. The method of claim 17,wherein the molecular environment comprises a reactant capable ofgenerating a product and wherein the reactant is capable of reactingwith the reactive moiety to modulate the reactive moiety between thefirst state and the second state.
 21. The method of claim 20, whereinthe reactant comprises H⁻, OH⁻, an acid, a base, an acetyl, a methyl, aphosphoryl, carboxyl, hydroxyl, or amine.
 22. The method of claim 17,wherein the molecular environment comprises an ion capable of pairingwith the charged moiety to modulate the charged moiety between the firststate and the second state.
 23. The method of claim 22, wherein the ioncomprises a member selected from the group consisting of ammonium,imidazolium, pryidinium, pyrrolidinium, phosphonium, sulfonium, and anycombination thereof.